Mitochondrial DNA: Methods and Protocols 1071629212, 9781071629215

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Methods in Molecular Biology 2615

Thomas Nicholls · Jay Uhler Maria Falkenberg Editors

Mitochondrial DNA Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-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.

Mitochondrial DNA Methods and Protocols

Edited by

Thomas J. Nicholls Wellcome Centre for Mitochondrial Research, Newcastle University, Newcastle upon Tyne, UK

Jay P. Uhler Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden

Maria Falkenberg Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden

Editors Thomas J. Nicholls Wellcome Centre for Mitochondrial Research Newcastle University Newcastle upon Tyne, UK

Jay P. Uhler Department of Medical Biochemistry and Cell Biology University of Gothenburg Gothenburg, Sweden

Maria Falkenberg Department of Medical Biochemistry and Cell Biology University of Gothenburg Gothenburg, Sweden

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2921-5 ISBN 978-1-0716-2922-2 (eBook) https://doi.org/10.1007/978-1-0716-2922-2 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 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 Mitochondria, the energy-generating and metabolic hubs of eukaryotic cells, possess their own genome called mitochondrial DNA or mtDNA. Mitochondrial genomes vary widely in size and structure between kingdoms, from small and compact molecules in mammals to massive, complex multimeric genomes in some plants, but are united by their central role in encoding components of the oxidative phosphorylation system. Human mtDNA was first discovered in the 1960s and is a multicopy genome found in the form of thousands of individual nucleoprotein complexes, called nucleoids, spread around the mitochondrial network. The copy number of human mtDNA is maintained by a DNA replication machinery of largely bacteriophage and bacterial origin, which operates independently of nuclear DNA replication and the cell cycle. Detailed genetic studies of mtDNA using classical molecular genetic techniques have long been hampered by the lack of a routine system to transform the mitochondria of human cells, although these systems exist for some model organisms, such as yeast. In vitro studies have also proven invaluable for the study of mitochondrial genetic processes. An inability to maintain sufficient copies of mtDNA per cell, or the accumulation of mutations or deletions in a subset of mtDNA molecules, is a cause of mitochondrial disease in humans. These rare but devastating metabolic diseases can manifest at any stage of life and can result from defects arising in the mtDNA itself, or alternatively from pathological variants in the nuclear genes that encode the mtDNA replication and expression machineries. In this volume of Methods in Molecular Biology: Mitochondrial DNA, now in its fourth edition, we aim to provide a broad range of protocols that can be used to study mtDNA on molecular, cellular, and whole organism levels, both in vivo and in vitro. The book is divided into six parts. Part I provides methods for the isolation of mitochondria and mtDNA from different species and cell types, which can be used as a starting point for further analysis. In Part II, protocols are presented for the visualization and quantification of mtDNA using different microscopy techniques. Part III contains methods for identifying and characterizing the mtDNA-interacting proteins of the mitochondrial nucleoid. Protocols for the analysis of mtDNA replication and repair are given in Part IV, utilizing reconstituted in vitro systems as well as using material isolated from cells and tissues. Modern techniques for the in vivo modification of mtDNA are presented in Part V. Finally, Part VI provides methods for the investigation of mtDNA in pathological states and for genetic diagnostics in cases of human mitochondrial disease. We hope that these methods will be useful and informative for researchers and clinicians from all career stages with an interest in mitochondrial DNA, from students to seasoned professors. We are grateful to all of the authors that have contributed to the volume for their hard work and for their willingness to share their experience and expertise with the wider scientific community. Newcastle upon Tyne, UK Gothenburg, Sweden Gothenburg, Sweden

Thomas J. Nicholls Jay P. Uhler Maria Falkenberg

v

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

PART I

PURIFICATION METHODS FOR MITOCHONDRIAL DNA

1 Isolation of Functional Mitochondria and Pure mtDNA from Murine Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dieu Hien Rozsivalova, Milica Popovic, Harshita Kaul, and Aleksandra Trifunovic 2 Isolating Mitochondria, Mitoplasts, and mtDNA from Cultured Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katja E. Menger and Thomas J. Nicholls 3 Coupling Differential Centrifugation with Exonuclease Treatment and Size Exclusion Chromatography (DIFSEC) for Purification of mtDNA from Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrew M. Shaw and Payam A. Gammage 4 Isolation and Quality Control of Yeast Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . Asli Aras Taskin, Daiana Nerina Moretti, F. Nora Vo¨gtle, and Chris Meisinger 5 Mitochondrial DNA Isolation from Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fre´de´rique Weber-Lotfi, Arnaud Fertet, Rokas Kubilinskas, and Cle´mentine Wallet, and Jose´ M. Gualberto

PART II

v xi

3

17

31 41

57

VISUALISING MTDNA

6 Visualize the Distribution and Dynamics of Mitochondrial DNA (mtDNA) Nucleoids with Multiple Labeling Strategies. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Xiangjun Di, Jinshan Qin, Yujie Sun, and Qian Peter Su 7 Visualization of mtDNA Using FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Xie Xie and Xuefeng Zhu 8 In Situ Analysis of Mitochondrial DNA Synthesis Using Metabolic Labeling Coupled to Fluorescence Microscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 John A. Smolka and Samantha C. Lewis 9 Measurement of Nucleoid Size Using STED Microscopy . . . . . . . . . . . . . . . . . . . . 107 Elisa Motori

PART III 10

MITOCHONDRIAL DNA INTERACTING PROTEINS

How to Quantify DNA Compaction by TFAM with Acoustic Force Spectroscopy and Total Internal Reflection Fluorescence Microscopy . . . . . . . . . 121 Martial Martucci, Louis Debar, Siet van den Wildenberg, and Geraldine Farge

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Contents

11

Assessing TFAM Binding to Human Mitochondrial DNA . . . . . . . . . . . . . . . . . . . 139 Takehiro Yasukawa and Dongchon Kang 12 Identification of Proximity Interactors of Mammalian Nucleoid Proteins by BioID . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Mari J. Aaltonen and Hana Antonicka 13 Localization of Mitochondrial Nucleoids by Transmission Electron Microscopy Using the Transgenic Expression of the Mitochondrial Helicase Twinkle and APEX2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 David Pla-Martı´n, Felix Babatz, and Astrid C. Schauss

PART IV 14 15 16

17

18

19

20 21

22

In Vitro Assays of TWINKLE Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jay P. Uhler, Ulrika Alexandersson, and Maria Falkenberg Rolling Circle Replication and Bypass of Damaged Nucleotides . . . . . . . . . . . . . . Josefin M. E. Forslund, Gorazd Stojkovicˇ, and Sjoerd Wanrooij Studying Mitochondrial Nucleic Acid Synthesis Utilizing Intact Isolated Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jelena Misic and Dusanka Milenkovic Functional Assessment of Mitochondrial DNA Maintenance by Depletion and Repopulation Using 2’,3’-Dideoxycytidine in Cultured Cells. . . . . . . . . . . . . Ga´bor Zsurka, Genevieve Trombly, Susanne Scho¨ler, Daniel Blei, and Wolfram S. Kunz Analysis of Mitochondrial DNA Replication by Two-Dimensional Agarose Gel Electrophoresis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steffi Goffart and Jaakko Pohjoism€ a ki Quantitative Analysis of Nucleoside Triphosphate Pools in Mouse Muscle Using Hydrophilic Interaction Liquid Chromatography Coupled with Tandem Mass Spectrometry Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sushma Sharma, Ziqing Kong, Shaodong Jia, Phong Tran, Anna Karin Nilsson, and Andrei Chabes Detection of UV-Induced Deletions in Mitochondrial DNA . . . . . . . . . . . . . . . . . Gabriele A. Fontana and Hailey L. Gahlon Determination of the Ribonucleotide Content of mtDNA Using Alkaline Gels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Choco Michael Gorospe, Bruno Marc¸al Repoleˆs, and Paulina H. Wanrooij 5′-End Mapping in Human Mitochondrial DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . Andranik Durgaryan and Anders R. Clausen

PART V 23

MITOCHONDRIAL DNA REPLICATION AND REPAIR 191 203

219

229

241

267

281

293

315

MODIFYING MITOCHONDRIAL DNA

Manipulation of Murine Mitochondrial DNA Heteroplasmy with mtZFNs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 Pavel A. Nash and Michal Minczuk

Contents

ix

Biolistic Transformation of Chlamydomonas reinhardtii and Saccharomyces cerevisiae Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345 Nathalie Bonnefoy and Claire Remacle 25 A Method for Precisely Identifying Modifications to Plant Mitochondrial Genomes by mitoTALENs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Tomohiko Kazama and Shin-ichi Arimura 24

PART VI

MITOCHONDRIAL DNA IN HUMAN DISEASE

26

Mitochondrial DNA Sequencing and Heteroplasmy Quantification by Next Generation Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrea Legati, Daniele Ghezzi, and Carlo Viscomi 27 Genomic Strategies in Mitochondrial Diagnostics. . . . . . . . . . . . . . . . . . . . . . . . . . . Dasha Deen, Charlotte L. Alston, Gavin Hudson, Robert W. Taylor, and Angela Pyle 28 Mitochondrial DNA Enrichment for Sensitive Next-Generation Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shilan Wu, Matthew J. Longley, Scott A. Lujan, Thomas A. Kunkel, and William C. Copeland 29 Single Cell Analysis of Mitochondrial DNA Deletions . . . . . . . . . . . . . . . . . . . . . . . Helen A. L. Tuppen, Amy K. Reeve, and Amy E. Vincent Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

381 397

427

443 465

Contributors MARI J. AALTONEN • Montreal Neurological Institute, McGill University, Montreal, QC, Canada; Department of Human Genetics, McGill University, Montreal, QC, Canada ULRIKA ALEXANDERSSON • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden CHARLOTTE L. ALSTON • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Newcastle University, Newcastle upon Tyne, UK; NHS Highly Specialised Services for Rare Mitochondrial Disorders, Royal Victoria Infirmary, Newcastle upon Tyne Hospitals NHS Foundation Trust, Newcastle upon Tyne, UK HANA ANTONICKA • Montreal Neurological Institute, McGill University, Montreal, QC, Canada; Department of Human Genetics, McGill University, Montreal, QC, Canada SHIN-ICHI ARIMURA • Graduate School of Agriculture and Life Sciences, University of Tokyo, Tokyo, Japan FELIX BABATZ • Cologne Cluster of Excellence on Cellular stress response in Aging-associated Disease, CECAD, University of Cologne, Cologne, Germany DANIEL BLEI • Division of Neurochemistry, Institute of Experimental Epileptology and Cognition Research, University of Bonn, Bonn, Germany NATHALIE BONNEFOY • Institute of Integrative Biology of the Cell (I2BC), CEA, CNRS, Univ Paris-Sud, Universite´ Paris-Saclay, Gif-sur-Yvette cedex, France ANDREI CHABES • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden ANDERS R. CLAUSEN • Institute of Biomedicine, University of Gothenburg, Gothenburg, Sweden WILLIAM C. COPELAND • Genome Integrity and Structural Biology Laboratory, Mitochondrial DNA Replication Group, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA LOUIS DEBAR • Universite´ Clermont Auvergne, CNRS, Laboratoire de Physique de Clermont, Clermont-Ferrand, France DASHA DEEN • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Newcastle University, Newcastle upon Tyne, UK XIANGJUN DI • School of Biomedical Engineering, Faculty of Engineering and Information Technology, University of Technology Sydney, Sydney, NSW, Australia ANDRANIK DURGARYAN • Institute of Biomedicine, University of Gothenburg, Gothenburg, Sweden MARIA FALKENBERG • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden GERALDINE FARGE • Universite´ Clermont Auvergne, CNRS, Laboratoire de Physique de Clermont, Clermont-Ferrand, France ARNAUD FERTET • Institut de Biologie Mole´culaire des Plantes, CNRS, Universite´ de Strasbourg, Strasbourg, France GABRIELE A. FONTANA • Department of Health Sciences and Technology, ETH Zurich, Zurich, Switzerland

xi

xii

Contributors

JOSEFIN M. E. FORSLUND • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden HAILEY L. GAHLON • Department of Health Sciences and Technology, ETH Zurich, Zurich, Switzerland PAYAM A. GAMMAGE • CRUK Beatson Institute, Glasgow, UK; Institute of Cancer Sciences, University of Glasgow, Glasgow, UK DANIELE GHEZZI • Unit of Medical Genetics and Neurogenetics, Fondazione IRCCS Istituto Neurologico Carlo Besta, Milan, Italy; Department of Pathophysiology and Transplantation, University of Milan, Milan, Italy; Lab of Neurogenetics and ` Mitochondrial Disorders, Fondazione IRCCS Istituto Neurologico Carlo Besta/Universita degli Studi di Milano, Milan, Italy STEFFI GOFFART • University of Eastern Finland, Department of Environmental and Biological Sciences, Joensuu, Finland CHOCO MICHAEL GOROSPE • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden JOSE´ M. GUALBERTO • Institut de Biologie Mole´culaire des Plantes, CNRS, Universite´ de Strasbourg, Strasbourg, France GAVIN HUDSON • Wellcome Centre for Mitochondrial Research, Biosciences Institute, Faculty of Medical Sciences, Newcastle University, Newcastle upon Tyne, UK SHAODONG JIA • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden DONGCHON KANG • Department of Clinical Chemistry and Laboratory Medicine, Graduate School of Medical Sciences, Kyushu University, Fukuoka, Japan HARSHITA KAUL • Cologne Excellence Cluster on Cellular Stress Responses in AgingAssociated Diseases (CECAD) and Institute for Mitochondrial Diseases and Aging, Medical Faculty, University of Cologne, Cologne, Germany TOMOHIKO KAZAMA • Faculty of Agriculture, Kyushu University, Fukuoka, Japan ZIQING KONG • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden ROKAS KUBILINSKAS • Institut de Biologie Mole´culaire des Plantes, CNRS, Universite´ de Strasbourg, Strasbourg, France THOMAS A. KUNKEL • Genome Integrity and Structural Biology Laboratory, DNA Replication Fidelity Group, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA WOLFRAM S. KUNZ • Division of Neurochemistry, Institute of Experimental Epileptology and Cognition Research, University of Bonn, Bonn, Germany; Department of Epileptology, University of Bonn, Bonn, Germany ANDREA LEGATI • Unit of Medical Genetics and Neurogenetics, Fondazione IRCCS Istituto Neurologico Carlo Besta, Milan, Italy SAMANTHA C. LEWIS • Department of Molecular and Cell Biology, University of California, Berkeley, CA, USA MATTHEW J. LONGLEY • Genome Integrity and Structural Biology Laboratory, Mitochondrial DNA Replication Group, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA SCOTT A. LUJAN • Genome Integrity and Structural Biology Laboratory, DNA Replication Fidelity Group, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA

Contributors

xiii

MARTIAL MARTUCCI • Universite´ Clermont Auvergne, CNRS, Laboratoire de Physique de Clermont, Clermont-Ferrand, France CHRIS MEISINGER • Institute of Biochemistry and Molecular Biology, ZBMZ, Faculty of Medicine, University of Freiburg, Freiburg, Germany; CIBSS – Centre for Integrative Biological Signalling Studies, University of Freiburg, Freiburg, Germany; BIOSS Centre for Biological Signalling Studies, University of Freiburg, Freiburg, Germany KATJA E. MENGER • Wellcome Centre for Mitochondrial Research, Biosciences Institute, The Medical School, Newcastle University, Newcastle upon Tyne, UK DUSANKA MILENKOVIC • Max Planck Institute for Biology of Ageing, Cologne, Germany MICHAL MINCZUK • Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, UK JELENA MISIC • Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden DAIANA NERINA MORETTI • Institute of Biochemistry and Molecular Biology, ZBMZ, Faculty of Medicine, University of Freiburg, Freiburg, Germany; Faculty of Biology, University of Freiburg, Freiburg, Germany ELISA MOTORI • Institute for Biochemistry, University of Cologne, Cologne, Germany; Cologne Excellence Cluster on Cellular Stress Responses in Aging-Associated Diseases (CECAD), University of Cologne, Cologne, Germany PAVEL A. NASH • Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, UK THOMAS J. NICHOLLS • Wellcome Centre for Mitochondrial Research, Biosciences Institute, The Medical School, Newcastle University, Newcastle upon Tyne, UK ANNA KARIN NILSSON • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden DAVID PLA-MARTI´N • Center for Physiology, Faculty of Medicine and University Hospital, University of Cologne, Cologne, Germany; Center for Molecular Medicine Cologne, CMMC, University of Cologne, Cologne, Germany JAAKKO POHJOISMA€ KI • University of Eastern Finland, Department of Environmental and Biological Sciences, Joensuu, Finland MILICA POPOVIC • Cologne Excellence Cluster on Cellular Stress Responses in AgingAssociated Diseases (CECAD) and Institute for Mitochondrial Diseases and Aging, Medical Faculty, University of Cologne, Cologne, Germany ANGELA PYLE • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Newcastle University, Newcastle upon Tyne, UK JINSHAN QIN • State Key Laboratory of Membrane Biology, Biomedical Pioneer Innovation Center (BIOPIC), School of Life Sciences, Peking University, Beijing, China AMY K. REEVE • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Framlington Place, Newcastle University, Newcastle upon Tyne, UK CLAIRE REMACLE • Genetics and physiology of microalgae, InBios/Phytosystems Research Unit, University of Liege, Liege, Belgium BRUNO MARC¸AL REPOLEˆS • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden DIEU HIEN ROZSIVALOVA • Cologne Excellence Cluster on Cellular Stress Responses in AgingAssociated Diseases (CECAD) and Institute for Mitochondrial Diseases and Aging, Medical Faculty, University of Cologne, Cologne, Germany

xiv

Contributors

ASTRID C. SCHAUSS • Cologne Cluster of Excellence on Cellular stress response in Agingassociated Disease, CECAD, University of Cologne, Cologne, Germany SUSANNE SCHO¨LER • Department of Epileptology, University of Bonn, Bonn, Germany SUSHMA SHARMA • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden ANDREW M. SHAW • CRUK Beatson Institute, Glasgow, UK JOHN A. SMOLKA • Department of Molecular and Cell Biology, University of California, Berkeley, CA, USA GORAZD STOJKOVICˇ • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden QIAN PETER SU • School of Biomedical Engineering, Faculty of Engineering and Information Technology, University of Technology Sydney, Sydney, NSW, Australia YUJIE SUN • State Key Laboratory of Membrane Biology, Biomedical Pioneer Innovation Center (BIOPIC), School of Life Sciences, Peking University, Beijing, China ASLI ARAS TASKIN • Institute of Biochemistry and Molecular Biology, ZBMZ, Faculty of Medicine, University of Freiburg, Freiburg, Germany; CIBSS – Centre for Integrative Biological Signalling Studies, University of Freiburg, Freiburg, Germany ROBERT W. TAYLOR • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Newcastle University, Newcastle upon Tyne, UK; NHS Highly Specialised Services for Rare Mitochondrial Disorders, Royal Victoria Infirmary, Newcastle upon Tyne Hospitals NHS Foundation Trust, Newcastle upon Tyne, UK PHONG TRAN • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden ALEKSANDRA TRIFUNOVIC • Cologne Excellence Cluster on Cellular Stress Responses in AgingAssociated Diseases (CECAD) and Institute for Mitochondrial Diseases and Aging, Medical Faculty, University of Cologne, Cologne, Germany GENEVIEVE TROMBLY • Division of Neurochemistry, Institute of Experimental Epileptology and Cognition Research, University of Bonn, Bonn, Germany HELEN A. L. TUPPEN • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Framlington Place, Newcastle University, Newcastle upon Tyne, UK JAY P. UHLER • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden SIET VAN DEN WILDENBERG • Universite´ Clermont Auvergne, CNRS, Laboratoire de Physique de Clermont, Clermont-Ferrand, France; Universite´ Clermont Auvergne, CNRS, IRD, Universite´ Jean Monnet Saint Etienne, LMV, Clermont-Ferrand, France AMY E. VINCENT • Wellcome Centre for Mitochondrial Research, Translational and Clinical Research Institute, Faculty of Medical Sciences, Framlington Place, Newcastle University, Newcastle upon Tyne, UK CARLO VISCOMI • Department of Biomedical Sciences, University of Padova, Padova, Italy F. NORA VO¨GTLE • CIBSS – Centre for Integrative Biological Signalling Studies, University of Freiburg, Freiburg, Germany; Center for Molecular Biology of Heidelberg University (ZMBH), Heidelberg, Germany CLE´MENTINE WALLET • Institut de Biologie Mole´culaire des Plantes, CNRS, Universite´ de Strasbourg, Strasbourg, France PAULINA H. WANROOIJ • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden

Contributors

xv

SJOERD WANROOIJ • Department of Medical Biochemistry and Biophysics, Umea˚ University, Umea˚, Sweden FRE´DE´RIQUE WEBER-LOTFI • Institut de Biologie Mole´culaire des Plantes, CNRS, Universite´ de Strasbourg, Strasbourg, France SHILAN WU • Genome Integrity and Structural Biology Laboratory, Mitochondrial DNA Replication Group, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA XIE XIE • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden TAKEHIRO YASUKAWA • Department of Molecular Pathogenesis, Juntendo University Graduate School of Medicine, Tokyo, Japan XUEFENG ZHU • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden GA´BOR ZSURKA • Division of Neurochemistry, Institute of Experimental Epileptology and Cognition Research, University of Bonn, Bonn, Germany; Department of Epileptology, University of Bonn, Bonn, Germany

Part I Purification Methods for Mitochondrial DNA

Chapter 1 Isolation of Functional Mitochondria and Pure mtDNA from Murine Tissues Dieu Hien Rozsivalova, Milica Popovic, Harshita Kaul, and Aleksandra Trifunovic Abstract Detailed analysis of mitochondrial function cannot be achieved without good quality preparations of isolated mitochondria. Ideally, the isolation protocol should be quick, while producing a reasonably pure pool of mitochondria that are still intact and coupled. Here, we describe a fast and simple method for the purification of mammalian mitochondria relying on isopycnic density gradient centrifugation. We describe specific steps that should be taken into consideration when functional mitochondria from different tissues should be isolated. This protocol is suitable for the analysis of many aspects of the organelle’s structure and function. Key words Mitochondria, Isopycnic density gradient centrifugation, Organelle isolation, Electron transport chain, Oxidative phosphorylation, Respirometry, Blue native PAGE

1

Introduction Mitochondria are organelles present in almost every eukaryotic cell. They can occupy up to 25% of the volume of the cytoplasm. Furthermore, mitochondria play a central role in cellular life and death. They are often called “the power plants” of the eukaryotic cell for being the main sites of ATP production needed to drive various cellular processes. For decades, mitochondria were cast in this limited, but essential role. This view has drastically changed over last decade, as novel roles for mitochondria as a central hub of metabolism and cellular signaling have been described [1, 2]. Beyond their role as powerhouses that generate ATP, mitochondria are essential for producing various anabolic precursors and signaling metabolites that exert a profound effect on different cellular processes ranging from cellular growth to tumorigenesis. It

Dieu Hien Rozsivalova, Milica Popovic and Harshita Kaul contributed equally with all other contributors. Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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has also become abundantly clear that mitochondria do not work in isolation, but rather as a communicative reticulum in which they rapidly change size and shape in response to cellular signals. Therefore, it does not come as a surprise that dysfunctional mitochondria and/or dysregulated communication between mitochondria and the rest of the cell underlie several pathophysiological conditions. Last but certainly not the least in this new era of mitochondrial biology is the emerging role of mitochondria as important players during microbial infection, cellular orchestrators of innate immune responses, and reservoirs of a multitude of inflammatory signals. This vast diversity of mitochondrial functions is now appreciated, not only by researchers who specifically study mitochondrial functions but also by others who come across mitochondria in their own research of other cellular components and processes. Although certain aspects of mitochondrial function could be addressed as a part of the interconnected network of the cell, detailed understanding of the organelle’s structure and physiology can only be addressed in isolated mitochondria. Indeed, mitochondrial research has vastly benefited from the possibility for preparations of organelles isolated from tissues based on differential centrifugation initially developed over 70 years ago [3]. Today, numerous protocols for the isolation of highly functional mitochondria are available [4–9]. The basic protocols for the isolation of crude, functional mitochondria further used in a wide range of studies most often involve two different centrifugation steps [4– 7]. In the first step after tissue homogenization, cellular and tissue debris, including membrane structures and nuclei, are pelleted on relatively low centrifugation speeds (600 × g – 1000 × g). In the second step, mitochondria are pelleted using higher centrifugation speeds (7000 × g – 10,000 × g). This is a crude preparation as these mitochondria are often pelleted together with other cellular components, e.g., ER, ribosomes, and microsomes, whose percentage can be diminished by subsequent washing steps. Over the years, many laboratories have developed multiple protocols for the isolation of highly functional mitochondria from different organisms, cell and tissue types. They most commonly differ in the type of monosaccharide used to preserve mitochondrial coupling in the isolation buffer (most often used are sucrose or mannitol), number of strokes used for the disruption of cells and tissues, or the centrifugation speeds and number of washing steps. For the majority of subsequent applications, these crude mitochondrial preps are well suited. These include functional assessments such as the measurement of respiration capacity and reactive oxygen species production; in organello replication, transcription or translation rates; organelle import efficiency; or the structural analyses of different mitochondrial structures, including respiratory chain complexes using different types of molecular biology techniques.

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In this chapter, we provide a basic mito-isolation protocol that we use for the isolation of functional, enriched, intact mitochondria. We further describe a minor adaptation of the same isolation protocol that allows its application in the isolation of these organelles from different samples and tissues, including adaptation of how to isolate different populations of mitochondria from the same sample (cytoplasmic and peri-lipid droplet mitochondria). We also address specific challenges that might be faced when using these methods with the ultimate goal to obtain organelles as functional as possible. Finally, we provide a basic protocol for mitochondrial DNA (mtDNA) purification from isolated mitochondria with additional recommendations for more elaborate protocols.

2 2.1

Materials Equipment

1. Potter S homogenizer with glass cylinders (for specific volumes, see respective protocols) including plunger (or see step 2.1.2 below). 2. (Alternatively!) Ground-in glass-on-glass Dounce homogeniser (loose (pestle A) and tight fit (pestle B), for manual homogenization). 3. Benchtop centrifuge with cooling option, corresponding rotors for 50 and 15 mL Falcon tubes, Falcon tubes as well as Eppendorf tubes, with capability for at least up to 11,000 × g. 4. pH meter. 5. Freezer at -80 °C.

2.2 Reagents and Consumables

1. Chemically resistant 1.5 mL, 15 mL, and 50 mL tubes and pipette tips. 2. 50 mL and 1.5 mL tubes, pipette tips, (optional) filters for sterile filtration. 3. Double-distilled water. 4. PBS (phosphate buffered saline buffer (10× Dulbecco’s) powder, A0965, AppliChem). 5. Sucrose (S7903, Sigma Aldrich). 6. KCl (Potassium chloride, A2939, AppliChem). 7. EDTA (ethylenediaminetetraacetic Aldrich).

acid,

E5134,

Sigma

8. TES Buffer (2-(Tris(hydroxymethyl)methylamino)ethane-1sulphonic acid, A1084, AppliChem). 9. Fatty acid-free BSA (bovine serum albumin, A6003, Sigma Aldrich).

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10. Subtilisin A (protease from Bacillus licheniformis, P5380, Sigma Aldrich). 11. KOH (potassium hydroxide pellets, Biochemica, A3871, AppliChem). 12. Protease inhibitor cocktail (S8830, Sigma Aldrich). 13. Mannitol (Mannitol Biochemica, A1903, AppliChem). 14. EGTA (Ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid, E4378, Sigma Aldrich). 15. HEPES (2-[4-(2-hydroxyethyl)-1-piperazinyl]ethanesulphonic acid, A3724, AppliChem). 16. SDS (sodium dodecyl sulfate, A2263, AppliChem). 17. NaCl (sodium chloride, A2942, AppliChem). 18. TRIS (Tris(hydroxymethyl)aminomethane, AppliChem).

A22644,

19. Proteinase K (endopeptidase K, 1.24568, Sigma Aldrich). 20. Potassium acetate (141479, AppliChem). 21. Isopropanol ((2-propanol, USP, BP, Ph. Eur.) pure, pharma grade, A0892, AppliChem). 22. Ethanol (ethanol absolute for analysis, A1613, AppliChem). 23. Ice. 24. Liquid nitrogen (N2). 2.3 Buffers and Solutions 2.3.1 Mitochondrial Isolation Buffers (MIB)

1. MIB1: 100 mM sucrose, 50 mM KCl, 1 mM EDTA, 20 mM TES, 0.2% fatty acid-free (FFA) BSA, pH 7.2 (see Notes 1 and 2). 2. MIB2: 100 mM sucrose, 50 mM KCl, 1 mM EDTA, 20 mM TES, pH 7.2 (see Notes 1 and 2). 3. MIB3: 100 mM sucrose, 50 mM KCl, 1 mM EDTA, 20 mM TES, pH 7.2, protease inhibitor cocktail (see Notes 2 and 3). 4. MIBb – MIB brain: 220 mM mannitol, 70.1 mM sucrose, 104 mM HEPES, 0.2% FFA BSA, pH 7.7 (see Note 2). 5. MIBad – MIB adipocyte (SHE-BSA): 50 mM sucrose, 5 mM HEPES, 2 mM EGTA, pH 7.2, 2% FFA BSA [10] (see Notes 2 and 4).

2.3.2 Sample Freezing Buffer

500 mM sucrose, 10 mM HEPES, pH 7.7 (see Note 2).

2.3.3 Mitochondria Lysis Buffer

MLB: 50 mM Tris/HCl pH 8.0, 100 mM NaCl, 2.5 mM EDTA, 0.5% SDS.

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7

Methods Investigating functional mitochondria that have been isolated from different tissues and from cultured cells offers unmatched advantages. This protocol illustrates a step-by-step procedure to obtain functional mitochondria with high yield from different tissues. The isolation procedures described here require 1–2 h, depending on the source of the organelles (Fig.1).

3.1 Isolation of Mitochondria from Adult Heart

Dissection of tissues followed by mincing

Supernantant subjected to high speed centrifugation to pellet the mitochondria

This protocol describes a method for isolation of crude mitochondria from mouse heart. The crude mitochondrial pellet contains peroxisomal, lysosomal, and ER contaminations – if necessary, mitochondrial fractions can be further purified with, e.g., density sucrose gradient [11] or enrichment with anti-TOM22 magnetic beads [12]. However, further purification results in lower mitochondrial yield as the risk of losing some amount of sample increases with every step [13]. Freshly isolated mitochondria are required for all functional analyses such as in organello assays (Fig. 2), measurements of oxygen consumption using the O2kRespirometer (Oroboros Instruments) (Fig. 3) or Seahorse XFe96 Analyzer (Agilent), or import assays. The pellet can also be stored at -80 °C after snap-freezing in liquid N2 for subsequent structural

Homogenization-tissue specific speed and number of strokes (refer to text for specifics)

Washing step -mitochondrial pellet resuspended in buffer without BSA

Low speed centrifugation to separate mitochondria and cell debris

Mitochondria in supernatant -pellet with cell debris discarded

Final high speed centrifugation

Pellet containing coupled mitochondria for downstream analysis

Fig. 1 Schematic representation of general mitochondrial isolation from various murine tissues. (Image created using BioRender)

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Fig. 2 Examples of downstream analyses done on isolated mitochondria. Mitochondrial native protein complexes and their supramolecular structures separated on a BN-PAGE gel, followed by (a) Coomassie staining; (b) in gel activity for NADH:ubiquinone oxidoreductase (Complex I of electron transport chain); (c) in gel activity for cytochrome c oxidase (Complex IV of electron transfer chain); (d) SDS-PAGE gel of radiolabeled in organello synthesized mitochondrial proteins

Fig. 3 Respiration of isolated mouse heart mitochondria. Blue curve signifies O2 concentration (Y axis on the left), while red curve shows the respiration rate (O2 flux per mg of protein, Y axis on the right). An Oroboros protocol for isolated mitochondria was followed (SUIT-008-O2_mt_D026), in summary: Isolated mitochondria were supplemented with substrates in saturating concentrations to observe maximal capacity of oxidative phosphorylation (OXPHOS). Afterwards, FCCP was titrated to uncouple OXPHOS and to observe maximal capacity of the electron transport chain (ETC). Rotenone and antimycin A inhibit ETC which allows correction for residual oxygen consumption. The chamber was partially opened two times to reoxygenate the system. Legend: 1mt - isolated mitochondria, 1P - pyruvate, 1 M - malate, 2D - ADP, 3G - glutamate, O - open chamber, C - closed chamber, 4S - succinate, 5U - titration of FCCP, 6Rot - rotenone, 7Ama - antimycin A

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studies including: isolation of mtDNA and mtRNA, SDS-PAGE, BN-PAGE (Fig. 2), immunoprecipitation, ribosomal gradient centrifugation, etc. Basic Method All steps should be performed at 4 °C as much as possible. Tubes, centrifuges, and homogenizer equipment should be precooled to 4 °C. MIB should be precooled to 4 °C and kept on ice throughout the procedure. 1. Prepare MIB1 by adding FFA BSA to MIB2 just before use and keep on ice. 2. Prepare subtilisin A (1 mg/mL) in MIB2 (see Note 5). 3. Isolate mouse heart, remove arteries and blood clots. 4. Weigh the tissue. 5. Wash out blood from the tissue in PBS or MIB1. 6. Cut the tissue into smaller pieces with precooled equipment and transfer into 5 mL MIB1 in a 50 mL Falcon tube. Keep on ice. 7. Add subtilisin A to the MIB1 just prior to homogenization (1 μL subtisilin/1 μg tissue). 8. Transfer into a 5 mL glass homogeniser tube. 9. Homogenize at 1000 rpm with approximately 20 long strokes (until the solution is homogeneous). Perform homogenization at 4 °C. 10. Transfer the solution into a 50 mL Falcon tube and fill up to 20 mL with MIB1. 11. Centrifuge at 8500 × g for 5 min at 4 °C. 12. Discard supernatant to remove floating fat and subtilisin. 13. Resuspend pellet in 25 mL MIB1 without subtilisin A with a Pasteur pipette or by vigorous shaking. 14. Centrifuge at 800 × g for 5 min (4 °C). 15. Collect the supernatant into a new 50 mL tube, store on ice. Be careful not to disturb and collect the pellet. 16. To increase the relative mitochondrial yield, repeat steps 11– 13. 17. Combine supernatants from steps 13 and 14. 18. Centrifuge at 8500 × g for 5 min (4 °C) to pellet mitochondria. 19. Resuspend the pellet in 600–800 mL MIB2 and transfer to 1.5 mL tubes. 20. Centrifuge at 8500 × g for 5 min (4 °C) to pellet mitochondria. 21. Discard supernatant.

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22. Resuspend the mitochondrial pellet in 50–200 μL MIB3 (for functional analyses, use MIB2 as the protease inhibitor cocktail may interfere with downstream procedures). The volume of MIB2 or MIB3 depends on the mitochondrial yield (see Notes 6 and 7). 23. When performing functional analyses, keep on ice for no longer than 2 h. For long-time storage, snap-freeze in liquid N2 and store at -80 °C. For high-resolution respirometry using the O2k-Respirometer on mitochondria isolated from heart (Fig. 3), we recommend mitochondrial preparation using a different isolation buffer (MIB1 in this case: 225 mM mannitol, 75 mM sucrose, 1 mM EGTA, pH 7, 2.5 mg/mL BSA), decreasing the number of strokes during homogenization and omitting the first high speed centrifugation step, as described in a protocol provided by Oroboros Instruments (bioblast.at). In this case, the protocol has been adjusted to make sure mitochondria stay intact as it is necessary for oxidative phosphorylation to be coupled. This basic isolation method provides adequate mitochondria for further analyses of native mitochondrial protein complexes with BN-PAGE and in gel activity assays (Fig. 2a–c). It is also possible to do in organelle assays, which are used to assess de novo protein synthesis using radioactive labelling, since the mitochondria are coupled and intact (Fig. 2d). These mitochondria could be further used for the isolation of pure mtDNA (see Subheading 3.8). 3.2 Isolation of Mitochondria from Neonatal Heart

When isolating mitochondria from the hearts of neonatal mice, the following needs to be taken into consideration: (i) the amount of tissue is much smaller, so the equipment, amount of buffer, and homogenization step need to be scaled down; (ii) instead of a Potter homogenizer, a glass-on-glass Dounce homogeniser is used for homogenization by hand; (iii) mitochondria are homogenized in 1 mL MIB instead of 5 mL, and all the centrifugation steps are done in 5 mL of buffer in 15 mL Falcon tubes instead of higher volumes of buffer in 50 mL Falcon tubes. The specific steps are as follows: 1. Dissect heart from neonatal mice and transfer to 1 mL of cold MIB1. 2. After dissection, wash off blood residues as indicated in the basic protocol (Subheading 3.1, steps 3–5), and transfer tissue to a 2 mL glass homogeniser. 3. Add subtilisin as (Subheading 3.1).

indicated

in

the

basic

protocol

4. Homogenize tissue by hand with 15 slow strokes using the A pestle (loose), then 10 strokes using the B pestle (tight).

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5. Transfer samples to 15 mL Falcon tubes and add up to 5 mL of MIB1. Continue with the basic protocol (Subheading 3.1, step 11) for adult heart with adjusted volumes (see Note 7). 3.3 Isolation of Mitochondria from Intestine

When isolating mitochondria from fragile tissues such as the gut epithelia of mice (see Note 8), perform homogenization using a glass-on-glass Dounce homogenizer using 10 strokes with pestle A in MIB1, without addition of subtilisin. It is very easy to overhomogenize the tissue, which would lead to breakage of the nucleus and release of DNA into the supernatant containing mitochondria. This increases the viscosity of the sample, making downstream analysis, such as BN-PAGE very difficult. To decrease viscosity, samples can be treated with nuclease such as Benzonase, or they can be strained through a syringe needle (0.40 × 0.20 mm).

3.4 Isolation of Mitochondria from Liver

For the isolation of mitochondria from liver, use the same buffer and centrifugation steps as for adult heart, without addition of subtilisin. Homogenization should be performed using a Potter homogeniser at 1200 rpm with 10 long strokes.

3.5 Isolation of Mitochondria from Brain

Brain tissue, composed of different cell types rich in mitochondria, has high fat content and very soft consistency. Therefore, a primary concern is overhomogenization of the tissue that can result in rupture of mitochondria. Due to this, the buffer used for mitochondrial isolation from brain contains mannitol instead of sucrose, while the centrifugation and homogenization steps are critical points in obtaining high yield. The following steps are adjusted in comparison to the basic protocol: 1. Isolate brain tissue, cut into smaller pieces, and transfer to 5 mL MIBb (see Note 9). 2. Transfer into a 5 mL glass cylinder and homogenize in a Potter homogenizer with 10 long strokes at 1000 rpm (see Note 10). 3. Transfer homogenate into new 50 mL Falcon tube. 4. Continue with the basic protocol using MIBb (Subheading 3.1, steps 14–21). 5. Finally, resuspend mitochondria in sample freezing buffer (see 2.3.2) and snap-freeze in liquid N2.

3.6 Isolation of Mitochondria from Skeletal Muscle

This isolation protocol is adjusted for adult mouse quadriceps. If processing less tissue, downscaling is recommended. The isolation buffer used is the same as for the basic protocol, for the adult heart. The following steps should be followed:

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1. Dissect muscle and separate fat tissue thoroughly. 2. Put freshly isolated muscle tissue into a petri dish with cold MIB1 and cut into smaller pieces. 3. Transfer minced tissue into a 5 mL glass cylinder for homogenization. 4. Add subtilisin and MIB1 to final volume of 5 mL. 5. Homogenize using a Potter homogeniser (1000 rpm 15–20 strokes). 6. Transfer homogenate into a 50 mL Falcon tube and add buffer to a final volume of 15 mL. 7. Spin at 1000 × g for 5 min and collect the supernatant in a new 50 mL Falcon tube (see Note 11). 8. Resuspend the remaining pellet in 15 mL and spin one more time at 1000 × g for 5 min to collect even more mitochondria. 9. Continue with the basic protocol (Subheading 3.1, step 17). 3.7 Isolation of Cytoplasmic and Perilipid Droplet Mitochondria from Adipocytes

Recent studies have revealed the existence of distinct populations of mitochondria within brown adipocytes (BAT). Mitochondria bound to lipid droplets (peridroplet mitochondria – PDMs) promote lipid droplet biogenesis and protect from lipotoxicity, while cytoplasmic mitochondria (CM) are specialized for fatty acid oxidation and energy production [10], allowing for simultaneous occurrence of lipid synthesis and breakdown. Though the presence of PDMs has been demonstrated only for BAT, previous studies have indicated the presence of lipid-associated mitochondria in other types of adipocytes as well. The protocol describes isolation of CMs and/or PDMs from various adipose tissues, including BAT, perigonadal white adipose tissue (pgWAT), and inguinal white adipose tissue (iWAT) (Fig. 4). 1. Harvest intrascapular BAT/ pgWAT/ iWAT and rinse twice in PBS. 2. Weigh and mince the tissues with fine scissors. 3. Suspend minced tissues in MIBad buffer (1 mL/100 mg tissue). 4. Homogenize with a glass-teflon electric laboratory tissue homogenizer (7 strokes), followed by further finer homogenization with a glass-glass 15 mL dounce homogenizer (13 strokes). 5. Transfer homogenate to a new 50 mL Falcon tube, wash the tube with MIBad to collect tissue remnants stuck to the walls of the homogenizing tube. 6. Centrifuge in a swinging bucket rotor at 900 × g for 10 min at 4 °C.

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iBAT

iWAT

eWAT Dissection of tissues followed by mincing

Homogenization-automated followed by manual

Washing followed by low speed centrifugation

To be repeated twice, pellet in falcon washed with buffer, the transferred to eppendorfs for final pelleting of mitochondria.

CMs

CMs

PDMs PDMs

Separartion of supernatant with CMs and scraping fat layer for PDMs

Vigorous resuspension

High speed centrifugation

Fig. 4 Schematic representation of isolation of mitochondria from adipose tissues. (Image created using BioRender)

7. Carefully remove the tube from the centrifuge and collect the supernatant with a long syringe, by plunging through the floating fat layer, along the sides of the Falcon tube, without disrupting the fat layer. This is the fraction that contains CMs. Immediately store the tube on ice. Make sure not to disrupt the debris pellet at the bottom of the tube. 8. Immediately, position the original Falcon tube horizontally on ice. 9. Carefully scrape the fat layer into a new ice-cold 50 mL tube and resuspend with the same amount of MIBad as the amount of the supernatant. Make sure to scrape any fat bits stuck to the tube’s wall with MIBad. This fraction contains the lipidassociated PDMs. Vigorously shake the Falcon tube to emulsify the fat layer and disrupt the PDMs from the fat droplets as much as possible (see Note 12). 10. Centrifuge the two tubes at 9000 × g, in a fixed angle centrifuge for 10 min at 4 °C. 11. Discard the supernatant.

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12. Wash with 1 mL MIBad and transfer the dissolved pellet to 1.5 mL Eppendorf tubes. Centrifuge at 9000 × g for 10 min at 4 °C. 13. Discard the supernatant. 14. Resuspend pellets with MIBad buffer w/o BSA. 15. Directly continue with further downstream analyses or snapfreeze in liquid N2 for long-term storage. 3.8 Basic Protocol for mtDNA Purification from Isolated Mitochondria

In most cases, analyses of mitochondrial genome do not require highly purified mtDNA, hence they are performed on DNA samples isolated from whole tissue. However, in certain cases, pure mtDNA is required and this is then obtained from isolated mitochondria. The purified mtDNA can be used for a variety of studies such as enzyme manipulations, Southern or South-western blotting (in certain cases, not always requiring pure mtDNA), cloning, some qPCR analyses, and high-resolution mtDNA mutation analysis. However, for the latter, we recommend using a protocol developed for the mtDNA mutation analysis [14, 15]. 1. Mix the mitochondria with 400 μL of freshly prepared MLB. 2. Incubate in a 50 °C water bath for up to 60 min or until the solution becomes clear. 3. Add 400 μL ice-cold isopropanol, mix by turning the tube up and down several times. Incubate on ice for 10 min. 4. Spin in a microcentrifuge at top speed (at least 13,000 × g) for 20 min at 4 °C. 5. Remove supernatant and wash pellet with 1 mL 70% ice-cold ethanol. 6. Spin in a microcentrifuge at top speed (at least 13,000 × g) for 20 min at 4 °C. 7. If necessary, repeat steps 4–6. 8. Remove the supernatant. Remove the trace amount of ethanol using a pipette tip. Air dry for 20–30 s (see Note 13). 9. Resuspend the DNA in 20 μL double-distilled water. For short-term storage, the mtDNA could be kept at 4 °C. For long-term storage keep the extracted DNA at -80 °C (see Note 14).

4

Notes 1. Since MIB1 and MIB2 have the same composition except for BSA, MIB2 can be prepared in advance and stored at 4 °C (optionally filter-sterilized). Then MIB1 can be prepared by adding fatty acid-free (FFA) BSA to MIB2 just before use.

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2. Adjust pH with KOH. 3. For the protease inhibitor cocktail, use the concentration according to the manufacturer’s instructions. 4. For freezing the mitochondrial pellet, use MIBad buffer without BSA. 5. Prepare the stock by dissolving subtilisin in MIB2, aliquot in 1.5–2 mL tubes and store at -20 °C. 6. For mitochondrial pellets from 80–90 μg heart, add 200 μL of MIB2 or MIB3. This can also depend on the method of choice used for protein concentration measurement. The goal is to have a concentration that fits approximately into the range of a standard curve used for determining protein concentration. If the mitochondria are too diluted, centrifuge at 8500 × g and resuspend in an appropriate amount of buffer. 7. Low mitochondrial yield can be a consequence of inefficient or overly harsh homogenization. When the homogenization is inefficient, mitochondria are pelleted with the cell debris. In this case, it is recommended to repeat resuspension of cell debris and follow up with low-speed centrifugation to collect even more mitochondria. If the tissue is overhomogenized and mitochondria are broken, adjust the number of strokes and/or homogenization speed. If the amount of tissue is too low, increasing the amount of tissue or downscaling the protocol is recommended. 8. Washing out the gut with PBS may cause detachment of the epithelium, so it is not recommended. Instead, gut debris can be removed in low-speed centrifugation steps. 9. Depending on the amount of tissue or area of the brain, downscaling is recommended (as described in Subheading 3.2). 10. In case of high fat content: Centrifuge at 8500 × g for 5 min to separate fat. Discard supernatant and continue with the basic protocol by resuspending the pellet in corresponding buffer. 11. In case of contamination of mitochondrial pellet with cell debris, adjusting the low-speed centrifugation is recommended. Due to the low speed, cell debris can sometimes detach from the pellet and contaminate the supernatant. Adjust the centrifugation speed (up to 1000 × g) and/or time of centrifugation (e.g., increase from 5 to 10 min). It is also possible to use cell strainers (e.g., from VWR, nylon, 100 μm pores, 732-2759) or gauze to filter cell debris. 12. Instability of the fat layer or unstable fixing of the Falcon tube can lead to contamination of the PDM layer with CMs. Make sure to immobilize the Falcon tube on ice as much as possible, so that there is minimum movement while scraping the fat layer.

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13. Do not completely dry the DNA. It may be difficult to dissolve if it is completely dried. 14. Repetitive freeze-thaw cycles of the purified mtDNA sample will cause breakage that could be detected as deleted molecules when subsequent long-range qPCR (lr-qPCR) is used. Therefore, for these kinds of analyses, only freshly purified mtDNA should be used. References 1. Spinelli JB, Haigis MC (2018) The multifaceted contributions of mitochondria to cellular metabolism. Nat Cell Biol 20(7):745–754 2. Bock FJ, Tait SWG (2020) Mitochondria as multifaceted regulators of cell death. Nat Rev Mol Cell Biol 21(2):85–100 3. Hogeboom GH, Schneider WC, Pallade GE (1948) Cytochemical studies of mammalian tissues; isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate material. J Biol Chem 172(2):619–635 4. Hartwig S, Kotzka J, Lehr S (2021) Isolation and quality control of functional mitochondria. Methods Mol Biol 2276:41–55 5. Caldeira DAF, de Oliveira DF, Cavalcanti-deAlbuquerque JP, Nascimento JHM, Zin WA, Maciel L (2021) Isolation of mitochondria from fresh mice lung tissue. Front Physiol 12: 748261 6. Lebiedzinska-Arciszewska M, Wojtczak L, Wieckowski MR (2021) An update on isolation of functional mitochondria from cells for bioenergetics studies. Methods Mol Biol 2310: 79–89 7. Leterme S, Michaud M, Jouhet J (2021) Isolation of mitochondria for lipid analysis. Methods Mol Biol 2295:337–349 8. Fernandez-Vizarra E, Ferrin G, Perez-MartosA, Fernandez-Silva P, Zeviani M, Enriquez JA (2010) Isolation of mitochondria for biogenetical studies: an update. Mitochondrion 10(3): 253–262 9. Meyer EH, Millar AH (2008) Isolation of mitochondria from plant cell culture. Methods Mol Biol 425:163–169

10. Benador IY, Veliova M, Mahdaviani K, Petcherski A, Wikstrom JD, Assali EA, AcinPerez R, Shum M, Oliveira MF, Cinti S, Sztalryd C, Barshop WD, Wohlschlegel JA, Corkey BE, Liesa M, Shirihai OS (2018) Mitochondria bound to lipid droplets have unique bioenergetics, composition, and dynamics that support lipid droplet expansion. Cell Metab 27(4):869–885 e6 11. Clayton DA, Shadel GS (2014) Purification of mitochondria by sucrose step density gradient centrifugation. Cold Spring Harb Protoc 2014(10):pdb prot080028 12. Franko A, Baris OR, Bergschneider E, Von Toerne C, Hauck SM, Aichler M, Walch AK, Wurst W, Wiesner RJ, Johnston ICD, De Angelis MH (2013) Efficient isolation of pure and functional mitochondria from mouse tissues using automated tissue disruption and enrichment with anti-TOM22 magnetic beads. PLoS One 8(12):e82392 13. Hartwig S, Feckler C, Lehr S, Wallbrecht K, Wolgast H, Mu¨ller-Wieland D, Kotzka J (2009) A critical comparison between two classical and a kit-based method for mitochondria isolation. Proteomics 9(11):3209–3214 14. Kauppila JHK, Bonekamp NA, Mourier A, Isokallio MA, Just A, Kauppila TES, Stewart JB, Larsson NG (2018) Base-excision repair deficiency alone or combined with increased oxidative stress does not increase mtDNA point mutations in mice. Nucleic Acids Res 46(13): 6642–6669 15. Isokallio MA, Stewart JB (2021) Highthroughput detection of mtDNA mutations leading to tRNA processing errors. Methods Mol Biol 2192:117–132

Chapter 2 Isolating Mitochondria, Mitoplasts, and mtDNA from Cultured Mammalian Cells Katja E. Menger and Thomas J. Nicholls Abstract Mitochondria are double membrane-bound eukaryotic organelles with roles in a range of cellular activities including energy conversion, apoptosis, cell signalling, and the biosynthesis of enzyme cofactors. Mitochondria contain their own genome, called mtDNA, which encodes subunits of the oxidative phosphorylation machinery as well as the rRNA and tRNA molecules required for their translation within mitochondria. The ability to isolate highly purified mitochondria from cells has been instrumental in a number of studies of mitochondrial function. Differential centrifugation is a long-established method for the isolation of mitochondria. Cells are subjected to osmotic swelling and disruption, followed by centrifugation in isotonic sucrose solutions to separate mitochondria from other cellular components. We present a method using this principle for the isolation of mitochondria from cultured mammalian cell lines. Mitochondria purified by this method can be further fractionated to investigate protein localization, or act as a starting point to purify mtDNA. Key words Mitochondria, Mitochondrial DNA, Mitochondrial isolation, Protein localization, DNA extraction

1

Introduction Mitochondria play an essential role in a variety of cellular processes, such as oxidative phosphorylation (OXPHOS), apoptosis, fatty acid oxidation, and the biogenesis of iron-sulphur clusters and heme. Mitochondria are bound by two membranes: the outer mitochondrial membrane (OMM) and the inner mitochondrial membrane (IMM). The OMM is permeable to small molecules, while the IMM is an impermeable barrier across which the passage of protons is directed during OXPHOS. Around 1100 proteins are known to localize to mitochondria [1–3], and the impermeable nature of the IMM necessitates the use of a specialized import machinery for these proteins [4]. Mitochondria contain their own genome, mtDNA, which is a circular dsDNA molecule of (in humans) 16,569 bp in length

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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[5, 6]. Mitochondrial DNA encodes 13 polypeptides, all of which are components of the OXPHOS machinery, as well as the two rRNA and 22 tRNA molecules required for translation of these 13 proteins within mitochondria. The function of mtDNA is therefore inseparable from cellular energy metabolism. The isolation of mitochondria is a prerequisite for many studies of mitochondrial function. Mitochondria remain bioenergetically active after being isolated, and the ability to measure mitochondrial function following the addition of metabolic substrates and inhibitors has been used as the basis for a number of seminal discoveries of mitochondrial function. The lack of a routine system to transform mammalian mitochondria in order to perform reverse genetics [7] also accentuates the necessity for robust methods to study mtDNA in situ. Differential centrifugation is a classical method for the isolation of mitochondria from cells and solid tissues [8]. Briefly, a suspension of cells is mechanically disrupted in order to rupture the plasma membrane, and an isotonic sucrose solution is added to prevent the mitochondria from swelling and disintegrating. Centrifugation at low velocity is used to pellet and remove nuclei, and then centrifugation at higher velocity is used to separate crude mitochondria from the cytosol. Gradient centrifugation can then be used to further purify mitochondria from other organelles. Mitoplasts (mitochondria with the OMM removed) can also be generated by treatment with digitonin, which selectively solubilizes the IMM [9]. The breakage of cells causes significant quantities of both genomic DNA and non-mitochondrial proteins to associate with, and therefore copurify with, mitochondria during centrifugation. These can be removed by treatment of the intact mitochondria with DNase and proteinase, which are inhibited prior to lysis of the mitochondria. An outline of the workflow is shown in Fig. 1. Differential centrifugation can be used to assign a mitochondrial or submitochondrial localization to proteins of interest, by retaining fractions of intermediate purification steps and using western blotting to detect the protein of interest alongside marker proteins of known submitochondrial localization. Alternatively, purified mitochondria can be used as a starting point to isolate mtDNA free of nuclear contamination. Mitochondria and mtDNA isolated in this way are suitable for use in a variety of downstream applications, including many methods contained in this volume. These methods include in organello transcription and replication, visualization using microscopy, protein localization, deep sequencing, qPCR, Southern blotting, and 2D agarose gel electrophoresis.

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Cytosol Nuclei

Collect and homogenise cells

Low speed centrifugation

Crude mitochondria

High speed centrifugation

DNase treatment

Digitonin

Mitochondria

Proteins

Isolated mitochondria Proteinase K

Sucrose gradient sedimentation

mtDNA

Isolated mitoplasts

Fig. 1 Workflow for mitochondrial isolation using differential centrifugation. Cells are collected and disrupted using a Dounce homogenizer. Nuclei and unbroken cells are pelleted at low speed, and the supernatant is spun at higher speed to pellet mitochondria and other smaller cellular components. These are treated with DNase to remove genomic DNA, and mitochondria are further purified using sucrose gradient sedimentation. These isolated mitochondria can then be subfractionated using digitonin and proteinase K treatments, or used to isolate mtDNA

We present a protocol for mitochondrial isolation that can be used for the purification of mitochondria, mitoplasts, and mtDNA. As a long-established method, the procedure has been based upon, and adapted from, previously reported methods for mitochondrial isolation [10–13]. Alternative methods for mitochondrial isolation, such as the use of magnetic microbeads [14], also produce highquality mitochondria and may be preferable if using a small number of cells as starting material. The level of purity of mitochondria, and therefore the number of steps required in the isolation procedure, can be adjusted according to the experimental objective. Highly purified mitochondria or mitoplasts may be desirable for some experiments, but come at the expense of a longer protocol, lower end yield, and the need for a larger quantity of starting material.

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Materials Buffers for mitochondrial isolation should be sterile-filtered and stored at 4 °C. Buffer stocks are made without BSA, DTT, PMSF, or proteinase inhibitors, which should be added freshly. Aliquots of buffer stock are taken on the day of the experiment, to which the fresh buffer components are added. The quantity of homogenization and 2.5 × isotonic buffer required will depend upon the starting mass of cells. The 1 × isotonic buffer is also the basis for the DNase and protease buffers, and so these three buffers can be made from a single stock solution.

2.1 Crude Mitochondrial Isolation

1. Glass Dounce homogenizer with tight-fitting pestle, e.g., Kimble 15 mL Dounce tissue grinder. 2. Protease inhibitor tablets (e.g., Pierce Protease Inhibitor Tablets or Roche cOmplete Protease Inhibitor Cocktail). On the day of the experiment, dissolve one tablet in 1 mL sterile water to create a 50 × solution for addition to buffers. 3. 1 M 1,4-dithiothreitol (DTT) solution. Dissolve 1.54 g of powdered DTT to 10 mL with sterile water, aliquot, and store at -20 °C. 4. 100 mM phenylmethylsulfonyl fluoride (PMSF). Weigh out 174 mg of PMSF and add isopropanol to 10 mL. Aliquot and store at -20 °C (see Note 1). 5. 2% (w/v) digitonin solution. Prepare freshly on the day of the experiment (see Note 2). 6. Homogenization buffer: 20 mM HEPES pH 8, 5 mM KCl, 1.5 mM MgCl2, 2 mM DTT, 1 mg/mL BSA, 1 mM PMSF, 1 × protease inhibitors. Mix 10 mL of 1 M HEPES pH 8, 2.5 mL of 1 M KCl, and 750 μL of 1 M MgCl2, make up to 479 mL with sterile water, and sterile filter. On the day of the experiment, add DTT (from 1 M stock), BSA (from 100 mg/ mL stock), PMSF (from 100 mM stock), and protease inhibitors (from 50 × stock) to an aliquot of buffer stock to achieve the correct final concentrations. 7. 2.5 × isotonic buffer: 525 mM mannitol, 175 mM sucrose, 20 mM HEPES pH 8, 5 mM EDTA, 1 mg/mL BSA, 1 mM PMSF, 1 × proteinase inhibitors. Dissolve 47.82 g mannitol and 29.95 g sucrose in sterile water, add 10 mL 1 M HEPES pH 8 and 5 mL 0.5 M EDTA, make up to 479 mL and sterile filter. On the day of the experiment, add DTT, BSA, PMSF, and protease inhibitors as above to an aliquot of buffer stock to achieve the correct final concentrations. 8. 1 × isotonic buffer: 210 mM mannitol, 70 mM sucrose, 20 mM HEPES pH 8, 2 mM EDTA, 1 mM PMSF, 2 mM DTT,

Isolating Mitochondria, Mitoplasts, and mtDNA from Cultured Mammalian Cells

21

1 × proteinase inhibitors. Dissolve 19.13 g mannitol and 11.98 g sucrose in sterile water, add 10 mL 1 M HEPES pH 8 and 2 mL 0.5 M EDTA, make up to 479 mL and sterile filter. On the day of the experiment, add DTT, PMSF, and protease inhibitors as above to an aliquot of buffer stock to achieve the correct final concentrations. 9. DNase buffer: 210 mannitol, 70 mM sucrose, 20 mM HEPES pH 8, 2 mM EDTA, 10 mM MgCl2, 1 mg/mL BSA, 1 mM PMSF, 2 mM DTT, 1 × proteinase inhibitors (see Note 3). 10. DNase I powder or solution, e.g., deoxyribonuclease I from bovine pancreas (Sigma-Aldrich). 2.2 Sucrose Gradient Centrifugation

1. Benchtop ultracentrifuge, Beckman Coulter Optima MAX-XP or similar. 2. Swinging-bucket ultracentrifuge rotor and fitting ultracentrifuge tubes. We use a Beckman Coulter MLS-50 rotor with Beckman Coulter 5 mL open-top thin-wall polypropylene ultracentrifuge tubes. 3. Gradient buffer: 10 mM HEPES pH 7.8, 5 mM EDTA, 2 mM DTT (see Note 4). 4. 1 M sucrose in gradient buffer. Dissolve 3.42 g sucrose in gradient buffer and make up to 10 mL. Leave on a roller until fully dissolved and then keep on ice. 5. 1.5 M sucrose in gradient buffer. Dissolve 5.13 g sucrose in gradient buffer and make up to 10 mL. Leave on a roller until fully dissolved and then keep on ice. 1. Proteinase buffer: 210 mM mannitol, 70 mM sucrose, 20 mM HEPES pH 8, 2 mM EDTA, 2 mM DTT (see Note 5).

2.3 Mitoplast Preparation and Protease Treatment

2. Proteinase K, purchase as solution or dissolve to 20 mg/mL.

2.4 Cell Fractionation for Protein Localization

1. 2 × western lysis buffer: 100 mM Tris–HCl (pH 7.4), 300 mM NaCl, 2 mM EDTA, 2% (v/v) Triton X-100, 2 × proteinase inhibitor cocktail (from tablets). Freeze as 500 μL aliquots. 2. Nitrocellulose or polyvinylidene difluoride (PVDF) membrane.

2.5 mtDNA Extraction

1. DNA lysis buffer: 50 mM NaCl, 10 mM Tris–HCl pH 8, 10 mM EDTA, 0.5% SDS, 0.2 mg/mL proteinase K. 2. Phenol:chlofoform:isoamylalcohol (25:24:1) solution. 3. 175 mM NaCl solution 4. Isopropanol. 5. TE: 10 mM Tris–HCl pH 8, 1 mM EDTA. 6. Rotary mixer capable of accommodating microcentrifuge tubes.

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Methods Unless stated otherwise, all steps should be carried out on ice, preferably in a cold room. All centrifugation steps are carried out at 4 °C.

3.1 Crude Mitochondrial Isolation

1. Collect cells into a tube by an appropriate method. The number of cells required will depend upon the level of purity required and the downstream application (see Note 6). Adherent cells can be detached by trypsinization or by using a cell scraper. After collection, pellet cells by centrifugation at 300 g for 5 min. 2. Resuspend cells in PBS to remove culture medium, combine cells of the same type into a single tube, and pellet again at 300 g for 5 min. 3. Weigh the cell pellet using a laboratory balance (see Note 7). 4. Resuspend the cell pellet in 7 volumes of homogenization buffer, and leave on ice for 10 min (see Note 8). 5. Transfer the cell suspension to a glass Dounce homogenizer and disrupt cells using 10 strokes of a tight-fitting pestle (see Note 9). Place a sample of the cell suspension onto a microscope slide and assess the degree of cell disruption under a light microscope (see Note 10). Aim for approximately 80–90% of cells disrupted. 6. Transfer the cell homogenate to a Falcon tube and add a two-thirds volume of 2.5 × isotonic buffer (see Note 11). If performing cell fractionation for protein localization, retain a sample at this stage as the whole cell extract. 7. Centrifuge the homogenate at 1600 g for 10 min at 4 °C. Collect the supernatant into a new tube. If performing cell fractionation for protein localization, retain a sample of the pellet from this step as the nuclear/unbroken cells fraction. 8. Repeat the centrifugation of the supernatant at 1600 g for 10 min at 4 °C. Collect the supernatant into a new tube. 9. Centrifuge the supernatant from step 8 at 10,000 g for 10 min at 4 °C and remove the supernatant. If performing cell fractionation for protein localization, retain a sample of the supernatant from this step as the cytosolic fraction. 10. Resuspend the pellet in 10 mL of DNase buffer. Add DNase I to a final concentration of 0.2 mg/mL and place on a roller at 4 °C for 1 h. 11. Add EDTA to a final concentration of 15 mM. Centrifuge at 10,000 g for 10 min at 4 °C to pellet crude mitochondria. Discard the supernatant.

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12. Resuspend the pellet in 5 mL of 1 × isotonic buffer and pellet by centrifugation at 10,000 g for 10 min at 4 °C. Discard the supernatant. 13. Wash mitochondria once more by resuspending the pellet in 5 mL of 1 × isotonic buffer and pelleting again by centrifugation at 10,000 g for 10 min at 4 °C. 3.2 Sucrose Gradient Centrifugation

1. Make two-step sucrose gradients by pipetting 1.5 M sucrose solution into an ultracentrifuge tube, and layering an equal volume of 1 M sucrose solution (both made up in gradient buffer) on top (see Note 12). For Beckman Coulter 5 mL thinwall polypropylene tubes, use 2.2 mL each of 1.5 M and 1 M sucrose solution. If starting from 1 to 2 g of cells, make two gradients, or for more starting material make four gradients. 2. Resuspend the mitochondrial pellet in 200 μL of 1 × isotonic buffer using wide bore tips (see Note 13). Carefully pipette the mitochondria onto the top of the gradient, dividing equally between the gradients. 3. Weigh the gradients using a balance to ensure equal mass. If necessary, adjust the mass by adding 1 × isotonic buffer. 4. Place gradients into buckets of a swing-out rotor and centrifuge at 120,000 g for 1 hour at 4 °C. This corresponds to 33,000 rpm for a Beckman Coulter MLS-50 rotor. 5. Following ultracentrifugation, the mitochondria form a band at the interface between the 1 M and 1.5 M sucrose layers. Carefully remove most of the upper 1 M sucrose layer, then collect the mitochondrial layer using a wide bore P1000 tip. 6. Add four volumes of gradient buffer with gentle mixing (see Note 14). 7. Pellet mitochondria by centrifugation at 10,000 g for 10 min at 4 °C. Discard supernatant. 8. Wash mitochondria once by resuspension in 5 mL of 1 × isotonic buffer followed by centrifugation at 10,000 g for 10 min at 4 °C. Discard the supernatant again.

3.3 Mitoplast Preparation and Protease Treatment

1. Resuspend mitochondrial pellet in 2 mL of 1 × isotonic buffer. If performing cell fractionation for protein localization, retain an aliquot at this stage as untreated purified mitochondria. If treating with multiple digitonin concentrations, divide mitochondria into equal volumes (see Note 15 and Fig. 2). Add digitonin to desired final concentration. For complete disruption of the outer mitochondrial membrane, add digitonin to 0.2%. Incubate on ice for 10 min (see Note 16). 2. Pellet mitoplasts by centrifugation at 15,000 g for 10 min at 4 °C, and discard supernatant.

Mitochondria Cytosol

Nuclei / unbroken cells

Katja E. Menger and Thomas J. Nicholls

Whole cell lysate

24

-

- 0.1% 0.2% Digitonin

-

+

-

+

-

+ Proteinase K [kDa] 15

H3

50

MRPS27

40

TOM20

15 75

AIF

50 1

2

3

4

5

6

7

8

9

Fig. 2 Mitochondrial protein localization using differential centrifugation and western blotting. K562 cells (approx. 3 × 108) were fractionated using differential centrifugation to isolate mitochondria. Digitonin was used to selectively solubilize the outer mitochondrial membrane (lanes 6–9), and proteinase K was used to degrade externally bound proteins (lanes 5, 7 and 9). H3 is used as a nuclear marker, and MRPS27 is used as a marker for the mitochondrial matrix. TOM20 (located in the outer mitochondrial membrane) is used as a control for the efficacy of proteinase K treatment. AIF (located in the intermembrane space) is used as a control for the solubilization of the outer mitochondrial membrane by digitonin. Any high-quality antibodies specific to proteins of known submitochondrial localization can be used for this purpose. The antibodies used in this figure are: H3 (Abcam Cat# ab1791, RRID: AB_302613, 1:50,000); MRPS27 (Proteintech Cat# 17280-1-AP, RRID: AB_2180510, 1:1000); TOM20 (Abcam Cat# ab78547, RRID:AB_2043078 1: 1000); and AIF (Cell Signaling Technology Cat# 4642, RRID:AB_2224542, 1: 1000)

3. Resuspend mitoplasts in 1 mL Proteinase buffer. If performing cell fractionation, divide mitoplasts into two equal volumes for -/+ proteinase samples. Add proteinase K to a final concentration of 25 μg/mL. Incubate at 37 °C for 30 min (see Note 17). 4. Add PMSF to a final concentration of 5 mM to inhibit the proteinase treatment. Pellet mitoplasts by centrifugation at 15,000 g for 10 min at 4 °C, and discard supernatant (see Note 18). 5. Resuspend mitoplasts in 1 mL of 1 × isotonic buffer and pellet again by centrifugation at 15,000 g for 10 min at 4 °C. Discard supernatant.

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25

6. Repeat step 5 to wash mitoplasts once more. 7. Snap freeze mitoplast pellets in liquid nitrogen and store at -80 °C, or proceed to cell fractionation for protein localization (Subheading 3.4) or mtDNA extraction (Subheading 3.5). 3.4 Cell Fractionation for Protein Localization

1. Mix an equal volume of each cellular fraction with 2 × western lysis buffer. Incubate on a rotary mixer for 30 min at 4 °C (see Note 19). 2. Centrifuge lysates at 11,000 g for 3 min at 4 °C, and collect the supernatant into a new tube. 3. Determine protein concentration in each fraction by BCA assay. 4. Load and run equal protein amounts on an SDS-PAGE gel. 5. Transfer proteins to a nitrocellulose or PVDF membrane and perform western blotting using antibodies specific to proteins of interest. Antibodies specific to marker proteins of different subcellular compartments are used as controls. Example marker proteins are shown in Fig. 2.

3.5 mtDNA Extraction

Mitochondrial DNA is typically purified by first performing a mitochondrial isolation, and then isolating DNA from this mitochondrial fraction. The purity of the mtDNA is therefore highly dependent upon the efficiency of removal of nuclear genomic DNA in previous steps (Fig. 3). A standard phenol-chloroform method is provided here for extraction of mtDNA from isolated mitochondria, but other methods of DNA extraction, such as column purification, can also be used (see Note 20). 1. Resuspend mitochondrial pellet in 1 mL of DNA lysis buffer and mix until thoroughly homogenized. 2. Add an equal volume of phenol:chloroform:isoamyl alcohol (25:24:1) and place on a rotating platform for 15 min at room temperature (see Note 21). 3. Centrifuge samples at 20,000 g for 15 min at 4 °C. Afterwards the sample should be clearly separated into two phases. 4. Pipette the aqueous (upper) phase into a new tube, being careful not to disturb the organic (lower) phase (see Note 22). 5. Add NaCl solution to a final concentration of 100 mM, then add one volume (equal to the recovered aqueous phase plus the salt solution) of isopropanol. Mix thoroughly by inverting the solution several times or by placing on a rotary mixer for a few minutes. 6. Incubate sample on ice for 20 min (see Note 23).

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Katja E. Menger and Thomas J. Nicholls mtDNA

a

b

BamHI 14,258

Ba

m

HI

III

nd

Hi

[bp] Human mtDNA 16,569 bp

Human mtDNA 16,569 bp

[bp]

HindIII 12,570 HindIII 11,680

HindIII 6,203 20,555 17,000 15,258 13,825 12,119 10,171

1000 100 10,000 8,000

10

6,000 5,000

1

4,000

0.1

Whole cells

log 10 mtDNA enrichment

c

-

+ + + + + + Mitochondria

DNase Sucrose gradient Proteinase K

3,000

2,000

1

2

3

4

Fig. 3 Isolation of mtDNA from purified mitochondria of HeLa cells. (a) Diagrams indicating the expected restriction products from restricting human mtDNA with BamHI or HindIII. (b) DNA extracted from sucrose gradient-purified mitochondria of HeLa cells was restricted with BamHI (lane 2) or HindIII (lane 3) and separated on 0.6% agarose and stained with ethidium bromide. Size markers are: 1 kb DNA ladder (New England Biolabs, lane 1) and GeneRuler High Range DNA ladder (Thermo Fisher Scientific, lane 4). (c) Enrichment of mtDNA during successive mitochondrial purification steps. During mitochondrial preparation from HeLa cells, samples were retained of whole cell extract, crude mitochondria (prior to DNase treatment), DNase-treated crude mitochondria, sucrose gradient-purified mitochondria, and proteinase K-treated mitochondria. DNA was extracted from each fraction, and the enrichment of mtDNA was determined using qPCR with a primer set specific to mtDNA (ND1) normalized to the amplification of a single-copy nuclear gene (B2M). The qPCR method was carried out according to the method described in [18]. The enrichment of mtDNA is shown as a log10-transformed value relative to the whole cell sample

7. Centrifuge sample at 20,000 g for 15 min at 4 °C to pellet DNA. Remove and discard the supernatant, being careful not to disturb the pellet. 8. Add 1 mL of 75% (v/v) ethanol in ultrapure water to wash the pellet. Centrifuge at 20,000 g for 5 min at 4 °C to pellet DNA again. 9. Carefully remove and discard the supernatant. Leave the tube with the lid open at room temperature for 30 min to air dry (see Note 24). 10. Resuspend the DNA pellet in TE buffer and store at 4 °C.

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4

27

Notes 1. PMSF is toxic. Weigh out in a fume hood wearing mask, gloves, and eye protection, and follow local disposal guidelines. PMSF may precipitate during storage and can be redissolved by gentle warming and mixing. 2. 2% (w/v) digitonin readily dissolves in water with warming to 37 °C, but will subsequently precipitate. Digitonin solutions should be made fresh on the day of the experiment. 3. The DNase buffer is the same as 1 × isotonic buffer with the addition of BSA and MgCl2 and can be made from the same stock on the day of the experiment. 4. Gradient buffer can be made on the day of the experiment. Alternatively, a stock of the buffer can be prepared without DTT, sterile filtered and stored at 4 °C, and then the DTT can be added on the day of the experiment. 5. The proteinase buffer is the same as 1 × isotonic buffer but without PMSF or protease inhibitor tablets, so can be made from the 1 × isotonic buffer stock on the day of the experiment. 6. We routinely collect cells from 2 to 4 confluent Nunc Square BioAssay Dishes (ThermoFisher cat. no. 166508), each of which has a culture area of 600 cm2, for preparation of highly purified mitochondria. For commonly used human cell types such as HeLa, this corresponds to approximately 1.5–3 × 108 cells. Larger cells, or cells growing to lower density, may require more dishes as starting material. 7. We find that some commonly used tubes for cell culture, especially 30 mL Universals, have a large variability in weight between tubes. It is therefore a good idea to weigh each individual tube before and after cell collection (rather than using one empty tube to zero the balance) in order to accurately measure the mass of cells collected in each tube. 8. The homogenization buffer is hypotonic relative to the cells, and so this incubation causes cells to swell before being disrupted by homogenization. 9. The amount of force applied during homogenization by different people is a potential source of experimental variation. Physical force is required to break the cells, and the resistance created by the vacuum under the pestle when pulling up the pestle should be noticeable. 10. The number of strokes of the Dounce homogenizer required to disrupt the cells varies by cell type. The use of ten strokes is generally suitable for cell lines such as HeLa and HEK293. The optimum number of strokes for other cell types can be

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determined experimentally, by assessing disruption using a light microscope and homogenizing further if necessary. 11. The addition of 2.5 × isotonic buffer at this step adjusts the sucrose and mannitol concentrations in the buffer to be isotonic with mitochondria, to prevent further swelling and disruption. 12. When layering the 1 M sucrose on top of the 1.5 M sucrose layer, pipette slowly and carefully to prevent the layers from mixing. The interface between the two layers should be visible as a sharp line. Holding the tube up to light while pipetting makes the interface easier to see. 13. The use of wide bore tips helps to maintain the integrity of mitochondria. Either wide bore tips can be purchased, or the end can be cut from standard pipette tips using a pair of clean scissors. 14. The addition of gradient buffer in this step is necessary to readjust the sucrose concentration to allow mitochondria to be pelleted in a standard benchtop centrifuge. 15. For protein localization, we typically divide the mitochondrial pellet into three equal fractions to give samples treated with 0%, 0.1%, or 0.2% digitonin. Each of these fractions is subsequently divided into two, to be incubated in the absence or presence of proteinase K, as shown in Fig. 2. 16. An alternative method for mitoplast preparation, the ‘swellcontract method’, has also worked well in our hands. Mitochondria are first allowed to swell in a hypotonic buffer, and then the inner membrane is caused to rapidly contract by the addition of ATP and magnesium [13, 15, 16]. Resuspend mitochondria in 20 mM potassium phosphate (pH 7.2) containing 0.02% BSA and incubate on ice for 20 min. Add MgCl2 and ATP to a final concentration of 1 mM each, incubate on ice for another 5 min, and then pellet and wash as described following digitonin treatment in Subheading 3.3. 17. Proteinase treatment can also be carried out on ice if there are particular concerns about the incubation of mitoplasts at 37 °C for the experiment being performed. 18. Following digitonin and proteinase K treatments, mitoplast pellets become looser and require careful pipetting to avoid aspirating the pellet. Using 1.5 mL microcentrifuge tubes (rather than 2 mL tubes) for these centrifugation steps helps to form more compact pellets that are easier to handle. 19. The whole cell extract and nuclear fractions contain a large amount of genomic DNA, which can cause problems with pipetting when loading and running the SDS-PAGE gel. To remove this genomic DNA, we frequently treat these two

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fractions with 250 U Benzonase for 30 min at 37 °C immediately prior to the addition of lysis buffer. 20. The choice of DNA extraction method is influenced by the downstream application. For example, phenol-chloroform extraction efficiently preserves the integrity of mtDNA replication intermediates [17], while column purification may be preferable for PCR or sequencing-based experiments. 21. A rotating mixer that turn tubes end over end should be used, as this provides much more efficient mixing than a rotary tube roller. 22. If the organic phase is disturbed using pipetting, centrifuge samples again to separate the phases and repeat removal of aqueous phase. 23. Following addition of salt and isopropanol, samples can be stored at -20 °C if they are not going to be used immediately. 24. It is essential that all ethanol from the washing step is removed. After pipetting off the majority of the wash buffer using a P1000, remove any residual remaining wash buffer using a P10 pipette before air drying.

Acknowledgments T.J.N. is supported by a Sir Henry Dale Fellowship jointly funded by Wellcome and the Royal Society (213464/Z/18/Z) and a Rosetrees and Stoneygate Trust Research Fellowship (M811). References 1. Rath S, Sharma R, Gupta R, Ast T, Chan C, Durham TJ, Goodman RP, Grabarek Z, Haas ME, Hung WHW et al (2021) MitoCarta3.0: an updated mitochondrial proteome now with sub-organelle localization and pathway annotations. Nucleic Acids Res 49:D1541–D1547 2. Smith AC, Robinson AJ (2016) MitoMiner v3.1, an update on the mitochondrial proteomics database. Nucleic Acids Res 44:D1258– D1261 3. Rhee HW, Zou P, Udeshi ND, Martell JD, Mootha VK, Carr SA, Ting AY (2013) Proteomic mapping of mitochondria in living cells via spatially restricted enzymatic tagging. Science 339:1328–1331 4. Wiedemann N, Pfanner N (2017) Mitochondrial machineries for protein import and assembly. Annu Rev Biochem 86:685–714 5. Nass MM, Nass S (1963) Intramitochondrial fibers with DNA characteristics. I. Fixation and

electron staining reactions. J Cell Biol 19:593– 611 6. Anderson S, Bankier AT, Barrell BG, de Bruijn MH, Coulson AR, Drouin J, Eperon IC, Nierlich DP, Roe BA, Sanger F et al (1981) Sequence and organization of the human mitochondrial genome. Nature 290:457–465 7. Gammage PA, Moraes CT, Minczuk M (2018) Mitochondrial genome engineering: the revolution may not be CRISPR-Ized. Trends Genet 34:101–110 8. Hogeboom GH, Schneider WC, Pallade GE (1948) Cytochemical studies of mammalian tissues; isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate material. J Biol Chem 172:619–635 9. Schnaitman C, Erwin VG, Greenawalt JW (1967) The submitochondrial localization of monoamine oxidase. An enzymatic marker for

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the outer membrane of rat liver mitochondria. J Cell Biol 32:719–735 10. Higuchi Y, Linn S (1995) Purification of all forms of HeLa cell mitochondrial DNA and assessment of damage to it caused by hydrogen peroxide treatment of mitochondria or cells. J Biol Chem 270:7950–7956 11. Yang MY, Bowmaker M, Reyes A, Vergani L, Angeli P, Gringeri E, Jacobs HT, Holt IJ (2002) Biased incorporation of ribonucleotides on the mitochondrial L-strand accounts for apparent strand-asymmetric DNA replication. Cell 111:495–505 12. Kornblum C, Nicholls TJ, Haack TB, Scholer S, Peeva V, Danhauser K, Hallmann K, Zsurka G, Rorbach J, Iuso A et al (2013) Loss-of-function mutations in MGME1 impair mtDNA replication and cause multisystemic mitochondrial disease. Nat Genet 45:214–219 13. Pallotti F, Lenaz G (2007) Isolation and subfractionation of mitochondria from animal cells and tissue culture lines. Methods Cell Biol 80: 3–44

14. Hornig-Do HT, Gunther G, Bust M, Lehnartz P, Bosio A, Wiesner RJ (2009) Isolation of functional pure mitochondria by superparamagnetic microbeads. Anal Biochem 389: 1–5 15. Parsons DF, Williams GR, Chance B (1966) Characteristics of isolated and purified preparations of the outer and inner membranes of mitochondria. Ann N Y Acad Sci 137:643–666 16. Murthy MS, Pande SV (1987) Malonyl-CoA binding site and the overt carnitine palmitoyltransferase activity reside on the opposite sides of the outer mitochondrial membrane. Proc Natl Acad Sci U S A 84:378–382 17. Reyes A, Yasukawa T, Cluett TJ, Holt IJ (2009) Analysis of mitochondrial DNA by two-dimensional agarose gel electrophoresis. Methods Mol Biol 554:15–35 18. Grady JP, Murphy JL, Blakely EL, Haller RG, Taylor RW, Turnbull DM, Tuppen HA (2014) Accurate measurement of mitochondrial DNA deletion level and copy number differences in human skeletal muscle. PLoS One 9:e114462

Chapter 3 Coupling Differential Centrifugation with Exonuclease Treatment and Size Exclusion Chromatography (DIFSEC) for Purification of mtDNA from Mammalian Cells Andrew M. Shaw and Payam A. Gammage Abstract Direct analysis of mtDNA using PCR-free methods is limited by the presence of persistent, contaminating nucleic acids originating from the nuclear genome, even following stringent mitochondrial isolations. Here we describe a method developed in our laboratory that couples existing, commercially available mtDNA isolation protocols with exonuclease treatment and size exclusion chromatography (DIFSEC). This protocol produces highly enriched mtDNA extracts from small-scale cell culture, with near-undetectable nuclear DNA contamination. Key words mtDNA, Purification, Size exclusion chromatography, Gel filtration, DNA

1

Introduction Owing to their bacterial ancestry, mitochondria contain a heritably and spatially separate genome (mtDNA). Mammalian mtDNA is a 16.5 kb, circular, multi-copy, and largely exonic genome that is anchored to the inner mitochondrial membrane (IMM) facing the matrix. It encodes mRNA for key protein subunits of the oxidative phosphorylation system (OXPHOS), alongside necessary ribosomal, and transfer RNAs required to translate these on mitochondrial ribosomes [1]. Like nuclear DNA (nDNA), mtDNA is susceptible to misprocessing or damage, and accumulation of mtDNA mutations is associated with several common human pathologies, including cancer [2]. Alongside mutagenic lesions, the impact of epigenetic modifications of mtDNA is also an area of interest, a long-standing and disputed question being whether nucleotide methylation is a potential epigenetic modifier of mtDNA expression [3, 4]. Understanding the contributions of such modifications to mitochondrial biology in normal and diseased states is of great interest to the field.

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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However, a limiting factor to any direct analyses of the mitochondrial genome is the persistent presence of contaminating nucleic acids, nucleotides, and oligonucleotides from the nuclear genome. While mtDNA is present at a far greater copy number than the nuclear genome, in almost all cellular contexts it contributes a minority of the total cellular DNA content (~0.1–0.5%). As such, the potential for nDNA contamination of mtDNA samples is significant. Historically, efforts to purify mtDNA have focussed on differential centrifugation, density gradients, and exonuclease treatments that allow for the specific degradation of linear contaminants, leaving the circular mtDNA intact [5–7]. However, these methods require large amounts of starting material, do not completely eliminate nDNA from samples, and do not address the abundance of free contaminating nucleotides or oligonucleotides liberated from nDNA through nuclease treatment of the sample. While these issues do not present a major hurdle to indirect analysis of mtDNA, using PCR-based methods for example, they are a major impediment to direct analysis of mtDNA using analytical chemistry and mass spectrometry approaches. Here we describe a novel technique, coupling differential centrifugation-based mtDNA purification protocols with exonuclease treatment and size exclusion chromatography (SEC) to obtain highly enriched mtDNA (DIFSEC) (Fig. 1a). Briefly, a commercially available kit provides a crude mitochondrial isolate through differential centrifugation, followed by deproteination of the sample and DNA precipitation to give an initial mtDNA extract. The precipitated mtDNA extract is treated with a linear DNA-specific exonuclease to degrade contaminating nDNA while sparing circular mtDNA. The resulting mixture of enriched mtDNA and low molecular weight (LMW) contaminants is then separated by SEC to produce highly purified mtDNA, with contaminating nDNA at the limit of PCR-based detection. This approach yields mtDNA purifications suitable for direct analytic methods using relatively small-scale cell cultures.

2 2.1

Materials Cell Culture

1. 100 mm2 culture dishes, 15 mL conical tubes. 2. 1× Dulbecco’s Modified Eagle Medium (DMEM), supplemented with 4.5 g/L D-glucose, Glutamax, and 100 mg/L sodium pyruvate. 3. 1× Phosphate buffered saline (PBS). 4. 10× Trypsin-ethylenediaminetetraacetic acid (trypsin-EDTA) solution. 5. Fetal calf serum. 6. U2OS osteosarcoma cell line (ATCC).

Coupling Differential Centrifugation with Exonuclease Treatment and Size. . .

A

Initial mtDNA enrichment

Exonuclease treatment

B Untreated ExoV

Size exclusion chromatography

LMW contaminants

33

ddPCR purity assessment

C **

mtDNA

+

-

-

-

+

+ +

-

-

- total DNA + mtDNA + SEC + ExoV

Fig. 1 DIFSEC – a method for mtDNA purification from mammalian cells. (a) Schematic highlighting the key steps in DIFSEC. An initial mtDNA enrichment via differential centrifugation and DNA precipitation is performed using a commercially available kit from Abcam (#ab65321). Following this, samples are treated with linear DNA-specific exonuclease V (RecBCD) to degrade contaminating nuclear DNA. Intact mtDNA is then separated from the degraded contaminants and the components of the exonuclease V reaction on a Superose 6 column, yielding a high purity mtDNA sample. Relative purity is confirmed via highly sensitive droplet digital PCR (ddPCR). (b) A representative chromatogram of 260 nm absorbance in mtDNA samples with and without exonuclease V treatment. Peaks corresponding to mtDNA and low molecular weight (LMW) contaminant elution volumes are indicated. (c) Fold enrichment of mtDNA (MT-ND1) relative to the nuclear genome (EIF2C1) for a column-free total DNA extract and a mtDNA enriched sample using the Abcam kit and DIFSEC. Nuclear DNA values in the DIFSEC samples were near the detection limit for ddPCR, which accounts for the large error observed. n = 3. One way ANOVA analyses were performed to determine statistical significance. Error bars indicate SD. **, p < 0.01

2.2 Mitochondrial DNA Isolation

1. Abcam mtDNA purification kit (ab65321). 2. 1× PBS (ice cold). 3. 70% Ethanol.

2.3 Column-Free Total DNA Isolation

1. Digestion buffer: 10 mM NaCl, 10 mM Tris–HCl, 10 mM EDTA, 0.5% SDS, pH 8.0. 2. Proteinase K 20 mg/mL.

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3. RNase A 20 mg/mL. 4. Sodium acetate 3 M (pH 5.2). 5. 70% and 98% ethanol (ice cold). 6. TE buffer. 2.4 Exonuclease V Treatment and SEC

1. Exonuclease V (RecBCD). 2. 10× NEBuffer 4 (New England BioLabs): 200 mM Trisacetate, 100 mM magnesium acetate, 500 mM potassium acetate, 10 mM DTT, pH 7.9. 3. Adenosine 5′-triphosphate (ATP). 4. Nuclease-free water. 5. AKTA explorer or equivalent column chromatography system with fraction collector. 6. Cytiva Superose 6 increase 10/300 GL column (#29091596). 7. TE Buffer.

2.5 Relative mtDNA Copy Number Determination by Droplet Digital PCR (ddPCR)

1. Bio-Rad Automated Droplet Generator. 2. Bio-Rad QX200 Droplet Reader. 3. Bio-Rad C1000 Touch™ Thermal Cycler with 96–Deep Well Reaction Module. 4. PCR Plate Sealer. 5. Bio-Rad DG32 Automated Droplet Generator Cartridges. 6. Bio-Rad Automated Droplet Generation Oil for Probes. 7. Bio-Rad ddPCR™ Droplet Reader Oil. 8. Bio-Rad Pipet Tips for the AutoDG™ System. 9. Bio-Rad ddPCR Supermix for Probes (No dUTP). 10. Bio-Rad ddPCR™ Gene Expression Assay: MT-ND1, Human. 11. Bio-Rad ddPCR™ Copy Number Assay: EIF2C1, Human.

2.6 General Laboratory Equipment

1. 1.5 mL centrifuge tubes. 2. Nanodrop 2000 spectrophotometer. 3. Qubit fluorometer. 4. Vortex. 5. Milligram-sensitive scales. 6. Benchtop centrifuge (for 1.5 mL and 15 mL tubes). 7. Cell scrapers. 8. Deep-well 96 well plates.

Coupling Differential Centrifugation with Exonuclease Treatment and Size. . .

3

35

Methods Carry out all procedures at room temperature unless stated otherwise.

3.1 Cell Culture and Preparation of Cell Pellets for Isolation of DNA

1. Plate cells onto 5 × 100 mm2 culture dishes and grow to 100% confluence. 2. Upon reaching confluence, wash cells two times in 10 mL of 1× PBS. 3. Scrape cells into 1 mL of ice-cold PBS and pool PBS/cell suspensions from each dish into a single 15 mL conical tube. 4. Transfer 1 mL of PBS/cell suspension into another tube for total DNA extraction. 5. Pellet the cells by centrifugation at 300 × g for 10 min at 4 °C. 6. Aspirate the supernatant and proceed to DNA isolation.

3.2 Column-Free Total DNA Isolation

1. Resuspend the pellet set aside for total DNA isolation in 500 μL of digestion buffer. Transfer to a 1.5 mL tube and add 2.5 μL proteinase K and RNase. Mix by pipetting or vortexing. 2. Incubate for 15 min at 55 °C. 3. Mix the sample again by pipetting or vortexing, then centrifuge at max speed for 2 min in a benchtop centrifuge. 4. Transfer the supernatant to a new 1.5 mL tube and discard the pellet. 5. Precipitate the protein and cell debris by adding 1/10 v/v 3 M sodium acetate (pH 5.2) for a final concentration of 300 mM. Pipette or invert to mix, and incubate at -20 °C for 10–15 min. 6. Centrifuge at max speed for 20 min at 4 °C. 7. Transfer the supernatant into a new 1.5 mL tube (see Note 1). 8. Precipitate the nucleic acids from the sample, by adding two volumes of ice-cold 98% ethanol (final ethanol percentage of 60–80%). 9. Invert to mix, and incubate at -20 °C for 10–15 min. 10. Centrifuge at max speed for 20 min at 4 °C. 11. Discard the supernatant. 12. Wash the pellet twice with 70% ethanol, dislodging the pellet each time. Centrifuge at max speed for 5–10 min to repellet the precipitate.

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13. Aspirate as much of the ethanol as possible without disturbing the pellet and allow to air dry. 14. Resuspend the DNA in 1× TE buffer and assess total DNA yield using a UV spectrophotometer. 3.3

mtDNA Isolation

3.4 Exonuclease V Treatment

Produce an initial mtDNA extract following the supplier’s protocols. Adjust volumes accordingly if using greater amounts of starting material than suggested in the protocol. 1. Pipette 50 μg of mtDNA prep into a new 1.5 mL tube. 2. Add the reaction components stated in Table 1 below: 3. Incubate at 37 °C for 4 h minimum (see Note 2).

3.5 Size Exclusion Chromatography

A key aim of this isocratic purification is to ensure maximal separation of the intact mtDNA from any contaminating nucleic acids, nucleosides, or oligonucleotides that have been produced by the exonuclease V reaction, as well as from the reaction components themselves. We recommend using a Superose 6 increase column 10/300 GL (see Note 4). This section will assume the user has an appropriate column chromatography system set up for use. 1. Rinse the column with 1.5 column volumes (CV) of doubly distilled and degassed H2O using a preconfigured or manual run. 2. Equilibrate the column with 1.5 CV of TE buffer with a preconfigured or manual run. 3. Load the 200 μL sample from 3.2 into an appropriate injection loop. 4. Run the sample on a preconfigured SEC and fraction collection program using the values detailed in Table 2 below. 5. Once the run is completed and fractions have been collected, note the fractions collected under the first UV 260 nm peak observed on the chromatogram (see Note 4 and Fig. 1b). These fractions will contain the purified mtDNA. Collect these fractions and proceed to copy number analysis. Table 1 Reaction components for Exonuclease V treatments Component

Amount (200 μL reaction)

NEBuffer 4 (10×)

20 μL

ATP (10 mM)

20 μL

Exonuclease V

10 μL (100 units)

Nuclease free H2O

Up to 200 μL final volume

Coupling Differential Centrifugation with Exonuclease Treatment and Size. . .

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Table 2 Details of SEC procedure Variable

Value

Flow rate (mL/min)

0.5

Column pressure limit (MPa)

3.00

Wavelength (nm)

260 (see Note 3) 280 215

Elution tube type

96 well

Eluate fraction size (mL)

0.5

Length of elution (column volumes)

1.2

Empty loop with volume (mL)

2.5

Table 3 Components of ddPCR reactions Component

Amount (per 20 μL reaction)

DNA (1 ng/μL)

1 μL

ddPCR supermix (2×)

10 μL

mtDNA primer/probe set (20×)

1 μL

Nuclear primer-probe set (20×)

1 μL

Nuclease free H2O

7 μL

A representative chromatogram of mtDNA samples applied to the Superose 6 column with and without exonuclease treatment is presented (Fig. 1b). A peak corresponding to low molecular weight (LMW) contaminants, including oligo- and mononucleotides liberated by exonuclease treatment, and a peak where mtDNA elutes in these conditions are indicated. 3.6 Assessment of Relative mtDNA Copy Number Using ddPCR

1. Thaw the ddPCR 2× super mix for probes on ice and place the refrigerated primer/probe sets on ice. 2. Dilute the mtDNA samples to 1 ng/μL (see Note 5). 3. Take a 96 well ddPCR plate and add 1 ng of mtDNA per sample replicate (see Notes 6 and 7). 4. Once thawed, make up a master mix containing the primer/ probe set and the supermix according to Table 3 below.

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Table 4 Cycling conditions for ddPCR reactions Step

Temperature (°C)

Time (min)

Ramp (°C/s)

1

95

10:00

2.5

2

94

0:30

2.5

3

55

1:00

2.5

4

Go to step 2 × 40

5

96

10:00

2.5

6

12

5:00

2.5

7

4

1

2.5

5. Add 19 μL of the master mix to each well of the 96 well plate that contains a sample. Include H2O containing wells as a non-template control (NTC). 6. Using a multichannel pipette, mix the samples by pipetting up and down, taking care not to generate bubbles (see Note 8). 7. Seal the plate and set up the droplet generator according to manufacturer’s instructions. 8. Once droplets have been generated, inspect the plate to ensure uniformity across all samples. Take note of any inconsistent wells. 9. Seal the droplet containing plate and run the ddPCR reaction on the thermo-cycler with conditions as in Table 4 below. 10. Once the PCR is completed, analyze the droplets on the droplet reader according to the manufacturer’s instructions. The relative purity of mtDNA from a total DNA sample, a mtDNA isolation sample, and mtDNA isolation samples treated with and without exonuclease V treatment and purified via SEC (DIFSEC) is shown in Fig. 1c. Note that the nuclear DNA signal in exonuclease V +DIFSEC purifications is near the detection limit for ddPCR. As such, these samples are expected to be essentially free from intact nuclear DNA contaminants and any oligonucleotides or mononucleotides liberated by exonuclease treatment.

4

Notes 1. Take care when removing the supernatant, ensuring that none of the pellet is carried into the next tube. 2. It may be advantageous to incubate Exonuclease V reactions overnight to ensure maximum activity against contaminating

Coupling Differential Centrifugation with Exonuclease Treatment and Size. . .

39

nucleic acids, but it should be noted that some loss of mtDNA may also be observed with these extended incubations. 3. The UV absorption peak for nucleic acids is 260 nm, and as such it is optimal to use a chromatography system that allows for detection at 260 nm. However, should the user only have access to a system with fixed UV detection at 280 nm (the absorption peak for protein), it is still possible to detect nucleic acids at this wavelength due to the protein depletion steps earlier in the protocol. We monitored three wavelengths to ensure the purity of the sample across the runs. 4. mtDNA is estimated to have a molecular weight of ~10 mDa, which is greater than the size exclusion limit of the Superose 6 column (fractionation range 5 kDa – 5 mDa). While mtDNA is not globular, and is topologically variable, its molecular weight is such that it will elute first within the void volume (peak at 260 nm). Degraded contaminants will enter the column resin, and elute much later towards the end of the run along with all of the exonuclease V reaction components. 5. Due to the multi-copy nature of mtDNA, the amount of DNA required to saturate the ddPCR reaction is relatively small. As such it may be difficult to find an amount of DNA that allows the user to reliably probe both mtDNA and nuclear DNA in the same reaction. If this is the case, probe both separately. Optimizing the amount of DNA input will be required on a cell-type by cell-type basis. 6. We recommend performing at least three replicates per sample to ensure technical reproducibility of the data. 7. Accuracy of DNA input is critical to achieving reliable ddPCR data. Accurate DNA quantitation for low concentration samples (e.g., Qubit) should be considered if data are not reproducible between technical replicates. 8. Bubbles can affect the droplet generation process and so should be avoided at all costs. If bubbles are an issue, we recommend preparing samples in a separate 96 well plate with several μL excess to allow for pipetting errors and prevent bubble formation when mixing. Once mixed, transfer 20 μL of sample to a fresh 96 well plate for droplet generation.

Acknowledgments The authors would like to thank Dr. Christopher Gray from Cytiva for helpful discussions regarding the use of SEC for these purposes. This work was supported by core funding to PG from CRUK BI (A_BICR_1920_Gammage). Figure 1a was created using BioRender.

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References 1. Gustafsson CM, Falkenberg M, Larsson NG (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160 Frezza C (2019) 2. Gammage PA, Mitochondrial DNA: the overlooked oncogenome? BMC Biol 17(1):53 3. Bicci I et al (2021) Oxford Nanopore sequencing-based protocol to detect CpG methylation in human mitochondrial DNA. bioRxiv 4. Patil V et al (2019) Human mitochondrial DNA is extensively methylated in a non-CpG context. Nucleic Acids Res 47(19):10072–10085

5. Devall M et al (2015) A comparison of mitochondrial DNA isolation methods in frozen post-mortem human brain tissue–applications for studies of mitochondrial genetics in brain disorders. BioTechniques 59(4):241–242, 244–6 6. Hao Z et al (2020) N6-deoxyadenosine methylation in mammalian mitochondrial DNA. Mol Cell 78(3):382–395.e8 7. Nakhle J et al (2020) Methods for simultaneous and quantitative isolation of mitochondrial DNA, nuclear DNA and RNA from mammalian cells. BioTechniques 69(6):436–442

Chapter 4 Isolation and Quality Control of Yeast Mitochondria Asli Aras Taskin, Daiana Nerina Moretti, F. Nora Vo¨gtle, and Chris Meisinger Abstract The isolation of organelles devoid of other cellular compartments is crucial for studying organellar proteomes and the localization of newly identified proteins, as well as for assessing specific organellar functions. Here, we describe a protocol for the isolation of crude and highly pure mitochondria from Saccharomyces cerevisiae and provide methods for testing the functional integrity of the isolated organelles. Key words Mitochondria, Saccharomyces cerevisiae, Differential centrifugation, Sucrose-gradient purification, Protein import, Osmotic swelling, Membrane potential

1

Introduction The yeast S. cerevisiae, with its short growth cycle, convenient lab handling, and well-characterized genome, has been an important model organism for numerous discoveries in life science including cell cycle regulation, autophagy, and protein trafficking [1, 2]. Budding yeast has also been an important and powerful tool for the analysis of the structure and function of the cellular powerhouse, mitochondria. Mitochondria are essential organelles in eukaryotes and not only provide the cell with energy, but harbor many important metabolic pathways and are a crucial checkpoint for the control of apoptosis [3]. Yeast has been intensively used to investigate basically all conserved organellar functions, including the structure and function of the respiratory chain, the biogenesis of the essential iron-sulphur cluster cofactors, the machineries for preprotein import and processing, the maintenance, expression, and inheritance of the mtDNA, organellar membrane dynamics via fusion and fission, and mitophagy [4–15]. Yeast also served as an important model for the analysis of the first comprehensive mitochondrial proteomes and subproteomes [16–22].

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Mitochondria isolated by early methods [23–25] were sufficiently pure to study specific mitochondrial functions, e.g., in organello protein import or respiratory activity. However, these fractions were significantly contaminated with other organellar compartments like endoplasmic reticulum, which is closely associated with, and difficult to separate from, mitochondrial membranes. Particularly, the analysis of mitochondrial protein composition or the analysis of dual-localized proteins revealed the need for methods yielding highly pure mitochondria largely devoid of other cellular compartments. An early method for purification of mitochondria was developed by Glick and Pon [26], which relied on separation of contaminant fractions from isolated mitochondria using a Nycodenz density gradient. Later, mitochondria isolation methods yielding mitochondria with much higher purity were established for the analysis of organellar proteomes and subproteomes and led to the identification of hundreds of novel mitochondrial proteins and protein modifications in yeast [16–22, 27, 28]. Here, we provide an updated protocol to obtain highly pure mitochondria, which are largely devoid of the ER fraction (as the most problematic contamination source (Fig. 1)). In addition, it is also highly important to test the structural and functional integrity of such highly purified mitochondria. Here, we have included protocols for two assays that allow quality control of the isolated organelles. First, the structural integrity of the outer mitochondrial membrane, which represents the most susceptible compartment to physical rupture or improper osmotic conditions, can be tested for

Fig. 1 Depletion of ER proteins in highly pure mitochondria compared to a standard isolation procedure. Immunoblot analysis of cellular fractions containing total yeast cells (Total), crude mitochondria (mito.), pure mitochondria, or the enriched microsomal/ER fraction (Microsom.). Samples were analyzed by SDS-PAGE (15%) and western blotting. Por1 and Tim23 are used as mitochondrial markers; Sec61 and Sss1 are used as ER markers

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43

Fig. 2 Protease accessibility upon osmotic swelling. Mitochondria were subjected to hypo-osmotic swelling (lanes 3–4). Where indicated, mitochondria (- swelling) and mitoplasts (+ swelling) were subsequently treated with Proteinase K (lanes 2 and 4). Samples were analyzed by SDSPAGE and immunodecoration. The outer membrane (OM) protein Tom22 and the OM form of Mcr1 are digested by Prot. K in mitochondria and mitoplasts. The intermembrane space (IMS) form of Mcr1 is only digested upon Prot. K treatment in mitoplasts. In contrast, the matrix proteins Mdh1 and Atp2 remain protected upon swelling

its ability to shield the intermembrane space against externally added proteases. As a control, hypoosmotic conditions are used for swelling of the tightly packed inner membrane and eventual disruption of the outer membrane (Fig. 2) [20]. Second, the preservation of membrane potential across the inner membrane during the isolation procedure can be monitored by import of the radiolabeled model precursor protein Su9-DHFR [22, 27, 29]. Su9-DHFR consists of the mitochondrial presequence of Neurospora crassa Fo-ATPase subunit 9 fused to the passenger protein dihydrofolate reductase (DHFR) from mouse [22, 29]. As the presequence directs the precursor protein to the mitochondrial matrix, the translocation across the inner membrane depends critically on an intact membrane potential. In this assay, we monitor time-dependent import of the Su9-DHFR precursor into a protease-protected compartment and maturation of the precursor by cleavage of its presequence upon entering into the matrix. The latter can be observed by a size shift after gel electrophoresis and also indicates the proper functioning of (proteolytic) enzymes in the matrix. As a control, the membrane potential is dissipated prior to the import reaction by addition of a mixture of the ionophore valinomycin, the complex III inhibitor antimycin A and the ATP synthase inhibitor oligomycin (AVO, see Subheading 2.4.2) to the reaction mix (Fig. 3).

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Fig. 3 Import of [35S]Su9-DHFR precursor protein into isolated mitochondria. Radiolabeled Su9-DHFR was incubated with wild-type mitochondria for the indicated times. Where indicated, the membrane potential (Δψ) was dissipated by addition of AVO (lanes 5 and 9) prior to the import reaction. Samples were treated with Proteinase K (Prot. K, lanes 6–9) to remove non-imported precursor protein. Samples were analyzed by SDS-PAGE and digital autoradiography. p precursor, m mature

2

Materials Prepare all buffers and solutions with distilled water and store buffers as listed below. All materials and labware for yeast growth need to be sterilized prior to use.

2.1

Yeast Culture

1. Medium for non-fermentative cell growth: YPG (1% [w/v] yeast extract (Difco), 2% [w/v] bacto peptone (Difco), 3% [w/v] glycerol, adjust pH to 4.9 with HCl). 2. Medium for fermentative cell growth: YPD (1% [w/v] yeast extract, 2% [w/v] bacto peptone, 2% [w/v] glucose, adjust pH to 4.9 with HCl). 3. Erlenmeyer flasks (for different culture volumes from 20 mL per flask for precultures up to 1.6–1.8 L per 5 L flask for main cultures). 4. Incubator shaker (e.g., New Brunswick, Innova 44/44R (with refrigeration)). 5. pH meter.

2.2 Isolation of Mitochondria

1. 1 M Tris–HCl, pH 7.4. Store at 4 °C. 2. 1 M Tris–H2SO4, pH 9.4. Store at 4 °C. 3. 1 M Dithiothreitol (DTT). Store in aliquots (e.g., 1 mL) at 20 °C. 4. 0.5 M Ethylenediaminetetraacetic acid (EDTA). Store at room temperature. 5. 2.4 M sorbitol. Keep at room temperature for direct use or store at 4 °C.

Yeast Mitochondria

45

6. 1 M MOPS-KOH, pH 7.2. Store at 4 °C. 7. 1 M K2HPO4. 8. 1 M KH2PO4. 9. 1 M KPi, pH 7.4: Titrate basic solution (1 M K2HPO4) with acidic solution (1 M KH2PO4) until pH 7.4. Store at 4 °C. 10. 0.2 M Phenylmethylsulfonylfluoride (PMSF). Prepare in isopropanol (stable for several months) and store at room temperature. 11. Protease Inhibitor Cocktail Tablets (e.g., cOmplete™ EDTAfree, Roche/Germany). 12. Zymolyase-20 T (Seikagaku Corporation, Tokyo, Japan). 13. Bovine serum albumin (BSA), fatty acid free, Sigma Aldrich, Germany. 14. Glass Teflon Dounce Homogenizer (20–60 mL volume). 15. Bradford reagent for protein estimation. 2.2.1

Working Solutions

1. DTT Buffer: 0.1 M Tris–H2SO4, pH 9.4, 10 mM DTT. Add DTT directly before use. DTT buffer should be heated/cooled to the same temperature as yeast growth temperature before use. 2. Zymolyase Buffer: 1.2 M sorbitol, 20 mM KPi, pH 7.4. Zymolyase buffer should be heated/cooled to the same temperature as yeast growth temperature before use. 3. Homogenization Buffer: 0.6 M sorbitol, 10 mM Tris–HCl, pH 7.4, 1 mM EDTA, 0.2% BSA, 1 mM PMSF. Add PMSF directly before use. PMSF is toxic, attention should be paid to avoid skin contact. Precool homogenization buffer to 4 °C. 4. SEM Buffer: 250 mM sucrose, 1 mM EDTA, 10 mM MOPS-KOH. Store at 4 °C.

2.3 Sucrose Gradient Purification of Isolated Mitochondria

1. Glass Teflon Homogenizer (5 mL volume). 2. Glass Pasteur pipettes (230 mm long), Carl Roth GmbH/ Germany). 3. Ultra-Clear™ Tubes (14 × 89 mm, Beckmann Coulter Inc./ USA). 4. Sorvall Discovery 90SE Ultracentrifuge. 5. Thermo Scientific™ TH-641 Swinging Rotor (including 6 titanium rotor buckets, bucket and rotor stand, closure tool for rotor). 5. EM Buffer: 1 mM EDTA, 10 mM MOPS-KOH. Store at 4 °C. 6. Sucrose stock solutions: prepare 60%/32%/23%/15% sucrose stock solutions in EM buffer. Store at 4 °C (stable for up to 1 month).

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2.4 Quality Control of Isolated Mitochondria

1. SEM Buffer: See Subheading 2.2.

2.4.1 Protease Accessibility Assay

3. 0.2 M Phenylmethylsulfonylfluoride Subheading 2.2.

2. EM Buffer: See Subheading 2.3. (PMSF):

See

4. Proteinase K (PK): 2.5 mg/mL stock in SEM buffer (used in final concentration of 10–50 μg/mL). Store in aliquots (e.g., 50 μL) at -20 °C. 5. Laemmli Buffer: 2% [w/v] SDS, 10% [v/v] glycerol, 0.02% [w/v] bromophenol blue, 62.5 mM Tris–HCl (pH 6.8), 1% [v/v] β-mercaptoethanol. Add β-mercaptoethanol directly before use. 6. SDS polyacrylamide gel electrophoresis standard equipment and materials. 7. Immunoblotting standard equipment and materials. 8. Chemiluminescence detection system (e.g., LAS4000 system, Fujifilm). 9. Image analysis software tools (e.g., Multigauge software (Fuji) and Photoshop (Adobe)). 2.4.2 In organello Import of Su9-DHFR into Isolated Mitochondria

1. [35S]-labeled Su9-DHFR precursor protein generated by in vitro transcription/translation using rabbit reticulocyte lysate system (Promega). See [30] for further details. 2. Import Buffer: 10 mM MOPS-KOH (pH 7.2), 3% [w/v] bovine serum albumin (BSA), 250 mM sucrose, 5 mM MgCl2, 80 mM KCl and 5 mM KPi. Store in aliquots (different volumes, e.g., 2 mL and 15 mL recommended) at -20 °C. 3. 0.2 M ATP. Prepare in H2O and titrate to pH 7.2 with KOH. Aliquot (e.g., 50 μL) and store at -20 °C. 4. 0.2 M NADH. Prepare freshly in SEM buffer. 5. AVO mix: 0.8 mM Antimycin A (inhibits electron transfer at complex III of the respiratory chain), 0.1 mM Valinomycin (potassium ionophore, uncouples oxidative phosphorylation), 2 mM Oligomycin (inhibitor of ATP Synthase). Prepare in ethanol. Store at -20 ° C. 6. Ethanol (p.a.). 7. Proteinase K: See Subheading 2.4.1. 8. SEM Buffer: See Subheading 2.2.1. 9. Laemmli Buffer: See Subheading 2.4.1. 10. SDS polyacrylamide gel electrophoresis standard equipment and materials. 11. Vacuum gel drying system.

Yeast Mitochondria

47

12. Phosphor screens and phosphor imaging system for digital autoradiography (e.g., Typhoon FLA7000 system). 13. Image analysis software tool for autoradiography (e.g., Image Quant™ software (GE Healthcare)).

3 3.1

Methods Yeast Culture

1. Inoculate a first preculture (20–50 mL medium) using yeast from a freshly grown plate. Grow cells overnight (16 h) at 30 ° C with vigorous shaking (temperature can vary depending on the strain background and experimental setup. In the following, standard growth conditions at 30 °C are used.). 2. Prepare a second preculture by diluting cells from the first preculture into 200 mL medium to a starting OD600nm of 0.2. Grow cells overnight (16 h) at 30 °C with vigorous shaking. 3. Prepare a main culture by diluting cells from the second preculture to 1.6–1.8 L medium in 5 L flasks to an OD600nm of 0.05. Incubate the main culture at 30 °C with vigorous shaking until an OD600nm of 1.5 is reached (see Note 1). The typical doubling time of wild type yeast cells grown on YPG medium at 30 °C is 3–4 h.

3.2 Isolation of Crude Mitochondria

All steps before cell homogenization should be carried out at room temperature (incubations should be carried out at the same temperature as cell growth). Homogenization and all subsequent steps are carried out on ice. 1. To harvest the cells, centrifuge cultures at 3000 g for 5 min at room temperature. Discard the supernatant. 2. Wash cell pellets with distilled water and centrifuge the cell suspension at 3000 g for 5 min at room temperature. Discard the supernatant. If the same strain was cultured in several flasks, combine cells to one sample. 3. Determine the pellet weight (the wet weight is used for calculation of the required volumes of DTT and Zymolyase buffers). 4. Gently resuspend cell pellet in pre-warmed DTT buffer (2 mL per gram wet weight). Incubate with shaking at 130 rpm at 30 ° C (growth temperature of main culture) for 20 min (see Note 2). 5. Centrifuge the cell suspension at 3000 g for 5 min at room temperature. Discard the supernatant. 6. Gently resuspend the pellet in 200 mL Zymolyase buffer (without enzyme) and centrifuge the cell suspension at 3000 g for 5 min at room temperature. Discard the supernatant.

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7. Resuspend cells in Zymolyase buffer (7 mL per gram wet weight). Keep 10 μL of this sample for the spheroplast test (label with ‘before zymolyase’). Calculate and weigh 3 mg zymolyase per gram of cells. Dissolve zymolyse in 1 mL Zymolyase buffer and add it to the cell suspension. Incubate for 30–45 min with shaking at 130 rpm and 30 °C (growth temperature of the main culture) (see Notes 2 and 3). 8. After the incubation, take 10 μL of sample to test for efficient zymolyase treatment. Label this sample with ‘after zymolyase’. Perform protoplast/spheroplast test by adding 1 mL of H2O and mixing of the sample by inversion. Assess the optical density of the samples or check for spheroplasts under the microscope (see Note 4). 9. Centrifuge the cell suspension at 3000 g for 5 min at room temperature. Discard the supernatant. 10. Carefully resuspend the pellet in 200 mL Zymolyase buffer (without enzyme) and centrifuge the cell suspension at 3000 g for 5 min at room temperature. Discard the supernatant. 11. Determine the pellet weight for calculation of the volume of homogenization buffer to be used. Cool down the centrifuge, centrifuge rotor and tubes to 4 °C for the following steps. 12. Resuspend pellet in ice-cold homogenization buffer (7 mL per gram wet weight) containing 1 mM PMSF (see Note 5). From this step onwards, work on ice and with ice-cold buffers. All following centrifugation steps are performed at 4 °C. 13. Homogenize samples with 20 strokes using a glass-Teflon dounce homogenizer. The cells are lysed by forcing them through the narrow space between glass and pestle. Use a potter size that fits to the volume (typically between 20 and 60 mL). The fit of pestle and glass determines the force and thereby the efficiency of cell breakage. Subject this homogenate to differential centrifugation as described in the following steps 14–19. 14. Centrifuge the homogenate at 1500 g for 5 min at 4 °C. Transfer the supernatant (containing the mitochondrial fraction) into a clean tube. Discard the pellet, which contains cell debris and nuclei. 15. Centrifuge the supernatant at 4000 g for 5 min at 4 °C. Transfer the supernatant (containing the mitochondrial fraction) into a clean tube. Discard the pellet. 16. To collect the mitochondrial fraction, centrifuge the supernatant at 17,000 g for 15 min at 4 °C. Discard the supernatant (see Note 6).

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17. Gently resuspend the mitochondrial pellet in 1 mL SEM buffer on ice, fill up to 20 mL with SEM buffer, and centrifuge at 4000 g for 5 min at 4 °C. Transfer the supernatant into a clean tube. Discard the pellet. 18. Centrifuge the supernatant at 17,000 g for 15 min at 4 °C. Discard the supernatant. 19. Resuspend the mitochondrial pellet in approximately 500 μL SEM buffer (see Note 7). 20. Determine the protein concentration by Bradford assay and adjust the concentration to 10 mg/mL with SEM buffer. 21. Aliquot mitochondria in desired volumes. Keep mitochondria on ice during this process. We aliquot the mitochondrial fraction into small fractions (typically 20–50 μL) to avoid freeze and thaw cycles. If isolation is followed by mitochondrial purification using a sucrose gradient (see Subheading 3.3), 3–5 mg aliquots are recommended. 22. Immediately after aliquoting, snap-freeze mitochondria in liquid nitrogen and store at -80 °C. 3.3 Isolation of Highly Purified Mitochondria

Carry out all steps on ice and with ice-cold buffers. Cool down all buffers, ultracentrifuge rotor and tubes before use. 1. For the sucrose gradient, the four different sucrose solutions are added from bottom to top into a Beckman ultracentrifuge tube with a Pasteur pipette. From the bottom start with 1.5 mL 60%, 3.5 mL 32%, 1.5 mL 23%, and 1.5 mL 15% sucrose (in EM buffer). Avoid bubble formation. 2. After finishing each sucrose gradient, check if the gradient interfaces are well-formed and store them at 4 °C. 3. Thaw crude mitochondria (from step 22 of Subheading 3.2) on ice and dilute to 5 mg/mL with SEM buffer (see Note 8). 4. Carefully homogenize the sample with 10 strokes using a small glass potter. The volume should not exceed 5 mL. 5. Load the homogenate on top of the sucrose gradient and centrifuge the samples at 100,000 g for 1 h at 2 °C. 6. Collect purified mitochondria from the interface between the 60% and 32% sucrose solutions and transfer into a fresh collection tube (see Notes 8 and 9). 7. Dilute the recovered mitochondria with two volumes of SEM buffer and centrifuge at 17,000 g for 15 min at 4 °C. 8. Determine the protein concentration by Bradford assay and adjust the concentration to 10 mg/mL with SEM buffer. 9. Aliquot mitochondria into desired volumes, snap-freeze in liquid nitrogen and store at -80 °C.

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3.4 Quality Control of Isolated Mitochondria 3.4.1 Protease Accessibility upon Osmotic Swelling

All steps are carried out on ice. 1. Prepare two samples by adding 50 μg mitochondria each into two 1.5 mL microcentrifuge tubes; label the first sample with – swelling and the second with + swelling. 2. Add 100 μL of SEM buffer and pellet mitochondria by centrifugation at 17,000 g for 10 min at 4 °C. Discard the supernatant. 3. Resuspend mitochondria in 400 μL of SEM buffer (-swelling) or 400 μL of EM buffer (+ swelling) by pipetting up and down 15 times using a combination of a yellow tip (10–200 μL volume) attached on top of a blue tip (100–1000 μL volume). 4. Incubate the samples on ice for 30 min and gently vortex every 10 min. 5. Divide the samples into two equal-sized aliquots using new microcentrifuge tubes. Label one of each new aliquot with +Prot. K and the other with -Prot. K. Add Proteinase K (final concentration 50 μg/mL) into the two +Prot. K samples and an equal volume of SEM buffer into the two -Prot. K samples. 6. Incubate samples on ice for 10 min. 7. Stop Proteinase K digestion by adding 8 μL of 0.2 M PMSF, and mix by gentle vortexing. 8. Incubate samples for 10 min on ice. 9. Pellet mitochondria by centrifugation at 17,000 g for 10 min at 4 °C. Discard the supernatant. 10. Wash mitochondrial pellets by the addition of 200 μL SEM buffer and pellet mitochondria by centrifugation at 17,000 g for 10 min at 4 °C. Discard the supernatant. 11. Resuspend pellets in 20 μL Laemmli buffer containing 1% β-Mercaptoethanol. 12. Incubate for 15 min at 65 °C with shaking at 14,000 rpm. 13. Analyze samples via SDS-PAGE followed by immunoblotting to assess Proteinase K accessibility of marker proteins of the different mitochondrial subcompartments (Fig. 2).

3.4.2 In organello Import of Su9-DHFR into Isolated Mitochondria

1. Label microcentrifuge tubes with 1, 2, 3 (three different timepoints, all samples without AVO (-AVO)) and tube 4 (last timepoint with AVO (+AVO)). 2. A typical single import reaction has a final volume of 100 μL and consists of 30–100 μg isolated mitochondria, 2 mM ATP, 4 mM NADH, the radiolabeled precursor, and import buffer (up to 100 μL). To dissipate the membrane potential, add 1 μL of AVO mix to tube 4. Add 1 μL of ethanol to the non-AVO

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samples as a control (see Notes 10 and 11). Mix gently by vortexing. 3. Keep all samples on ice. 4. To equilibrate the samples at the temperature of the import reaction, incubate samples at 25 °C for 3 min. 5. Add rabbit reticulocyte lysate containing [35S]-Su9-DHFR precursor protein (1–10% (v/v)) to each reaction tube (see Note 12). Mix gently with mild vortexing. 6. Incubate import reactions 1, 2, 3, and 4 at 25 °C for 2, 6, 18, and 18 min, respectively (see Note 13). 7. To terminate the import reaction, add 1 μL of AVO mix, mix gently by vortexing and place samples on ice. 8. Split samples into two equally sized aliquots by pipetting 50 μL of each sample into a new microcentrifuge tube (1–4: no Proteinase K treatment; 5–8: Proteinase K treatment). 9. Add 1 μL of SEM buffer to samples 1, 2, 3, and 4 and 1 μL of Proteinase K (final concentration 50 μg/mL) to samples 5, 6, 7, and 8. Mix gently by mild vortexing. Proteinase K digests non-imported precursors proteins. 10. Incubate samples on ice for 10 min. 11. Terminate proteinase K digestion by adding 1 μL of 0.2 M PMSF into all samples (tubes 1–8). Mix samples gently by mild vortexing. 12. Incubate samples on ice for 10 min. 13. Pellet mitochondria by centrifugation at 17,000 g for 10 min at 4 °C. Discard the supernatant (see Note 14). 14. Wash mitochondrial pellets by the addition of 200 μL SEM buffer and pellet mitochondria by centrifugation at 17,000 g for 10 min at 4 °C. Discard the supernatant. 15. Resuspend the pellets in 20 μL of Laemmli buffer containing 1% β-Mercaptoethanol. 16. Incubate at 65 °C for 15 min with shaking at 14,000 rpm. 17. Analyze samples by SDS-PAGE and digital autoradiography.

4

Notes 1. The doubling time of yeast cells depends on several factors (e.g., genotype, growth medium, temperature). Therefore, growth behavior of each strain has to be tested before mitochondrial isolation. The average yield of isolated mitochondria from 8 L yeast culture grown on respiratory medium (e.g.,

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YPGlycerol) to an OD600 of 1.5 is between 60 and 80 mg (protein amount). 2. The temperature of DTT and zymolyase incubations is the same as the growth temperature of the main cultures. Zymolyase treatment efficiency varies greatly at different temperatures, and the incubation time should be shortened (e.g., at 37 °C) or extended (e.g., at 19 °C) accordingly. 3. If mitochondria are isolated from yeast grown to a higher OD than 2, zymolyase digestion requires longer incubation than 30–45 min (up to 1 h). Alternatively, higher concentration of zymolyase (4–6 mg per g wet weight) can be applied. 4. We conclude that the cell wall is digested efficiently by zymolyase treatment, when the OD600nm of the ‘after zymolyase’ sample is reduced to one third of the OD600nm of the ‘before zymolyase’ sample. Usually, this is visible by eye when the ‘after zymolyase’ sample becomes clear and the ‘before zymolyase’ sample remains turbid. 5. For broader protease inhibition, add one protease inhibitor cocktail tablet (cOmplete™ EDTA-free) per 50 mL homogenization buffer prior to the homogenization procedure [21]. 6. This supernatant can be used to obtain cytosolic and microsomal fractions of yeast cells by ultracentrifugation at 100,000 g for 1 h at 4 °C (P, microsomal fraction; SN, cytosolic) [27]. 7. The volume of SEM buffer used for suspension of the final mitochondrial pellet depends on the starting material. We recommend 150 μL SEM buffer per gram of cells. 8. On average, one third of the protein amount of crude mitochondrial fractions consists of other contaminating cellular compartments and is therefore removed in the sucrosegradient purification. We usually start with 10 mg crude mitochondria and recover approximately 6–7 mg purified mitochondria from the 32/60% sucrose-gradient interface (see Fig. 4). 9. Approximately one third of the sample is typically present at the 15% and 23% sucrose interface (containing other cellular compartments and residual mitochondria) [27, 28]. 10. The 1 μL 100% ethanol is added into the – AVO samples to ensure equal sample treatment. 11. To test if the import of the precursor protein depends on the mitochondrial membrane potential, AVO mix is added to one sample (typically corresponding to the longest import time point) to dissipate the membrane potential across the inner membrane before the start of the import reaction.

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Fig. 4 Highly pure mitochondria are obtained from the 32/60% interface of the sucrose gradient (visible as a brownish ring)

12. We recommend the addition of 1–10% (v/v) reticulocyte lysate containing the radiolabeled precursor protein. 13. In organello import assays are usually carried out for up to 60 min. In the case of longer kinetics, we usually add an ATP-regenerating system into our import reaction mix, which consists of 5 mM creatine phosphate and 100 μg/mL creatine kinase. The import kinetics depend on several factors (e.g., import temperature, quality of mitochondria) and has to be established for each precursor protein individually. 14. The supernatant contains radioactively labeled (non-imported) precursor proteins and has to be collected and treated as radioactive waste.

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Acknowledgments Our work is supported by the Deutsche Forschungsgemeinschaft (DFG) under Germany’s Excellence Strategy (CIBSS – EXC-2189 – Project ID 390939984 to C.M., F.N.V. and A.A. T.), the RTG 2202/278002225 (to C.M.), RTG 2206/ 423813989 (to C.M. and F.N.V.), the SFB 1381 (Project-ID 403222702, to C.M. and F.N.V.), and the Emmy-Noether program (to F.N.V). References 1. Hartwell LH (1974) Saccharomyces cerevisiae cell cycle. Bacteriol Rev 38:164–198 2. Tsukada M, Ohsumi Y (1993) Isolation and characterization of autophagy defective mutants of Saccharomyces cerevisiae. FEBS Lett 333:169–174 3. Nunnari J, Suomalainen A (2012) Mitochondria: in sickness and in health. Cell 148:1145– 1159 4. Rutter J, Hughes AL (2015) Power(2): the power of yeast genetics applied to the powerhouse of the cell. Trends Endocrinol Metab 26: 59–68 5. Neupert W, Herrmann JM (2007) Translocation of proteins into mitochondria. Annu Rev Biochem 76:723–749 6. Shadel GS (1999) Yeast as a model for human mtDNA replication. Am J Hum Genet 65: 1230–1237 7. Mick DU, Fox TD, Rehling P (2011) Inventory control: cytochrome c oxidase assembly regulates mitochondrial translation. Nat Rev Mol Cell Biol 12:14–20 8. Sch€agger H, Pfeiffer K (2000) Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J 19:1777–1783 9. Lill R (2009) Function and biogenesis of ironsulphur proteins. Nature 460:831–838 10. Schmidt O, Pfanner N, Meisinger C (2010) Mitochondrial protein import: from proteomics to functional mechanisms. Nat Rev Mol Cell Biol 11:655–667 11. Wiedemann N, Pfanner N (2017) Mitochondrial machineries for protein import and assembly. Annu Rev Biochem 86:685–714 12. Poveda-Huertes D, Mulica P, Vo¨gtle FN (2017) The versatility of the mitochondrial presequence processing machinery: cleavage, quality control and turnover. Cell Tissue Res 367:73–81

13. Westermann B (2014) Mitochondrial inheritance in yeast. Biochim Biophys Acta 7:1039– 1046 14. Hoppins S, Lackner L, Nunnari J (2007) The machines that divide and fuse mitochondria. Annu Rev Biochem 76:751–780 15. Eisenberg T, Bu¨ttner S, Kroemer G et al (2007) The mitochondrial pathway in yeast apoptosis. Apoptosis 12:1011–1023 16. Sickmann A, Reinders J, Wagner Y et al (2003) The proteome of Saccharomyces cerevisiae mitochondria. Proc Natl Acad Sci U S A 100: 13207–13212 17. Zahedi RP, Sickmann BAM et al (2006) Proteomic analysis of the yeast mitochondrial outer membrane reveals accumulation of a subclass of preproteins. Mol Biol Cell 17:1436– 1450 18. Reinders J, Zahedi RP, Pfanner N et al (2006) Toward the complete yeast mitochondrial proteome: multidimensional separation techniques for mitochondrial proteomics. J Proteome Res 5:1543–1554 19. Vo¨gtle FN, Wortelkamp S, Zahedi RP et al (2009) Global analysis of the mitochondrial N-Proteome identifies a processing peptidase critical for protein stability. Cell 139:428–439 20. Vo¨gtle FN, Burkhart JM, Rao S et al (2012) Intermembrane space proteome of yeast mitochondria. Mol Cell Proteomics 11:1840–1852 21. Vo¨gtle FN, Burkhart JM, Gonczarowska-Jorge H et al (2017) Landscape of submitochondrial protein distribution. Nat Commun 8:290 22. Schmidt O, Harbauer AB, Rao S et al (2011) Regulation of mitochondrial protein import by cytosolic kinases. Cell 144:227–239 23. Daum G, Bo¨hni PC, Schatz G (1982) Import of proteins into mitochondria. Cytochrome b2 and cytochrome c peroxidase are located in the intermembrane space of yeast mitochondria. J Biol Chem 257:13028–13033

Yeast Mitochondria 24. Hartl FU, Ostermann J, Guiard B et al (1987) Successive translocation into and out of the mitochondrial matrix: targeting of proteins to the intermembrane space by a bipartite signal peptide. Cell 51:1027–1037 25. Zinser E, Daum G (1995) Isolation and biochemical characterization of organelles from the yeast, Saccharomyces cerevisiae. Yeast 11: 493–536 26. Glick BS, Pon LA (1995) Isolation of highly purified mitochondria from Saccharomyces cerevisiae. Methods Enzymol 260:213–223 27. Meisinger C, Sommer T, Pfanner N (2000) Purification of Saccharomyces cerevisiae

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mitochondria devoid of microsomal and cytosolic contaminations. Anal Biochem 287:339– 342 28. Meisinger C, Pfanner N, Truscott KN (2006) Isolation of yeast mitochondria. Meth Mol Biol 313:33–39 29. Walter C, Gonczarowska-Jorge H, Sickmann A et al (2018) Advanced tools for the analysis of protein phosphorylation in yeast mitochondria. Anal Biochem 554:23–27 30. Stojanovski D, Pfanner N, Wiedemann N (2007) Import of proteins into mitochondria. Meth Cell Biol 80:783–806

Chapter 5 Mitochondrial DNA Isolation from Plants Fre´de´rique Weber-Lotfi, Arnaud Fertet, Rokas Kubilinskas, Cle´mentine Wallet, and Jose´ M. Gualberto Abstract For most eukaryotes, sequencing and assembly of the mitochondrial DNA (mtDNA) is possible by starting the analysis from total cellular DNA, but the exploration of the mtDNA of plants is more challenging because of the low copy number, limited sequence conservation, and complex structure of the mtDNA. The very large size of the nuclear genome of many plant species and the very high ploidy of the plastidial genome further complicate the analysis, sequencing, and assembly of plant mitochondrial genomes. An enrichment of mtDNA is therefore necessary. For this, plant mitochondria are purified prior to mtDNA extraction and purification. The relative enrichment in mtDNA can be assessed by qPCR, while the absolute enrichment can be deduced from the proportion of NGS reads mapping to each of the three genomes of the plant cell. Here we present methods for mitochondrial purification and mtDNA extraction applied to different plant species and tissues, and compare the mtDNA enrichment obtained with the different procedures. Key words Mitochondria, mtDNA, Plants, Arabidopsis, Lettuce, qPCR, NGS sequencing

1

Introduction Mitochondria possess their own genome, the mtDNA, that vary in size, gene content, and organization between different organisms. In mammals, the mtDNA is a small circular and compact molecule [1], of a size of 16 kb in humans. In contrast, the mtDNA of higher plants varies extremely in size and in structural organization, even between closely related species. In most angiosperms, mtDNA can be mapped and represented as a single circular chromosome, of a size ranging between 200 and 700 kb. Yet, in certain species the mtDNA can be much bigger, up to 11.3 Mb in Silene conica, and can be encoded in multiple chromosomes [2]. However, despite their large sizes, plant mitogenomes only code for a limited number of protein genes and for several structural RNAs. Therefore, these genomes are mainly constituted by noncoding sequences that are not conserved and can represent up to 90% of the mtDNA, according to the species. An additional characteristic of plant

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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mitochondrial genomes is that they are rich in repeated sequences, which can be up to several tens of kilobases in size and are not conserved among species [3]. Recombination involving repeats leads to alternative configurations of the mtDNA and its distribution among interconvertible sub-genomic molecules, in a highly dynamic structure. Contrarily to mammals where the ploidy of the mtDNA is very high, in plants it is relatively low, with less than one copy per mitochondria [4]. The number of mtDNA copies per cell also varies according to the developmental stage and the cell type [5, 6]. All these characteristics of the plant mtDNA severely complicate the sequencing and assembly of plant mitogenomes, which nowadays is often the final goal of mtDNA purification. Furthermore, fragments of the mtDNA named NUMTs, for “nuclear mitochondrial DNA“, can be found in the nuclear chromosomes [7]. For instance, in Arabidopsis accession Col-0, there is a very large insertion in chromosome 2 comprising almost all mtDNA sequences [8]. These need to be taken into account in sequence assembly projects of plant mitogenomes. At present, the mtDNA of less than 300 land plant species have been sequenced and assembled. This number should rapidly increase with the widespread use of long-read sequencing technologies that facilitate genome assembly. But mtDNA sequencing can be difficult for species with very large nuclear genomes, which is often the case in plants. While the model plant Arabidopsis (Arabidopsis thaliana) has a small nuclear genome of just 135 Mbp, common lettuce (Lactuca sativa) has a genome as large as 2.5 Gbp, and wheat (Triticum aestivum) a genome of 14.5 Gbp. In those cases, the proportion of nuclear reads will be hundreds of times higher than the number of mitochondrial reads. Another important factor to take into account is the ploidy of the nuclear genome, which in species that undergo endoreplication, such as Arabidopsis, varies according to the type and age of the tissue [4, 9]. This means that in these plants it is important to extract the mtDNA from tissues where the proportion of mtDNA is the highest, such as young seedlings or inflorescences. Taking all of these considerations into account, to facilitate mtDNA sequencing and analysis, the best is to purify the mtDNA before sequencing. This involves the purification of mitochondria and extraction/purification of the mtDNA from the enriched mitochondrial fraction. The protocol for mitochondrial purification depends on the nature of the plant material. The hardness of the tissue will determine the procedure used for plant grinding (mortar and pestle, juice extractor, blender), the sugar concentration of the grinding buffer needs to be adapted to the water content of the tissue (to maintain osmotic pressure and avoid mitochondria lysis),

Plant mtDNA Purification Grinding

2 possible stages for DNase I treatment

Lysis of cells

Low speed centrifugation(s)

Remove of cell debris, chloroplasts and nuclei

High speed centrifugation

Sedimentation of crude mitochondria

Purification on Percoll gradient

Density purification of intact mitochondria

Washes

Elimination of Percoll

Centrifugation

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Sedimentation of purified mitochondria

Mitochondrial pellet

Fig. 1 Schematic procedure for plant mitochondrial purification

and the use of green material will lead to a greater contamination with chloroplastic DNA, as compared to etiolated tissues. Schematically, a plant mitochondrial purification consists of: grinding to lyse the rigid wall of the plant cells; low speed centrifugation(s) to remove cell debris, nuclei, and chloroplasts; high speed centrifugation to sediment mitochondria; purification on Percoll density gradients to separate intact mitochondria from broken ones and from plastids; washes to eliminate residual Percoll (Fig. 1). To eliminate nuclear and plastidial DNAs adhering to the outer mitochondrial membrane, a DNase treatment can be performed just before the Percoll gradient or at the end of the mitochondrial purification. Finally, to extract and purify the DNA from mitochondria, different methods exploiting different detergents for the lysis of mitochondria can be used, and the enrichment in mtDNA can be measured by qPCR or by the analysis of the proportion of NGS reads mapping to the mitochondrial genome. Here we describe the purification of mitochondria from two different plant tissues, Arabidopsis green seedlings and lettuce (Lactuca sativa) etiolated seedlings. We also describe a crude partial purification of the mtDNA that can be suitable for the analysis of single Arabidopsis plants. Different DNA extraction procedures are proposed and the enrichment in mtDNA is illustrated, both by the relative enrichment determined by qPCR and by the absolute coverage calculated from Illumina sequencing.

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Materials Plant Culture

2.1.1 In Vitro Culture of Arabidopsis Seedlings

1. Arabidopsis seeds, conserved desiccated at 15 °C in the dark. 2. Murashige & Skoog culture medium including vitamins and MES (M0255 from Duchefa-Biochemie, Haarlem, The Netherlands). 3. Culture medium: 0.5 × M0255 media (250 mg/L), 1% (w/v) sucrose, 0.8% (w/v) agar. Adjust to pH 5.8 with KOH before autoclaving the media. 4. Large Petri dishes (120 × 120 mm square plates). 5. Surgical Tape, 1.25 cm width (3 M™ Micropore™, Neuss, Germany). 6. Seed sterilizing solution: 70% (v/v) ethanol, 0.05% Triton X-100. 7. 70% ethanol and 90% (v/v) ethanol solutions. 8. Tabletop microcentrifuge (e.g., Eppendorf 5418R with standard rotor). 9. 1.5 mL microcentrifuge tubes. 10. Growth chamber or cabinet with controlled light and temperature. 11. Laminar air flow cabinet. 12. Sterile wooden toothpicks.

2.1.2 Culture of Arabidopsis Plants on Soil

1. 7 × 7 × 6.4 cm pots. 2. Soil mix. 3. Growth chamber or cabinet with controlled light and temperature.

2.1.3 Culture of Lettuce Seedlings on Blotting Paper

1. Lettuce seeds, conserved desiccated at 15 °C in the dark. 2. Sterile distilled deionized water. 3. Sterilized T-300 blotting paper disks (; 80 mm, All Paper b.v., Didam, Netherlands). 4. Petri dishes, high profile (; 90 mm × 50 mm, Dominique Dutcher, ref. 999283). 5. 1.5 mL microcentrifuge tubes. 6. 70% (v/v) ethanol. 7. Tabletop microcentrifuge (e.g., Eppendorf 5418R with standard rotor). 8. 1% NaOH. 9. Parafilm.

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10. Growth chamber or cabinet with controlled light and temperature. 11. Laminar air flow cabinet. 2.2 Extraction of Mitochondria 2.2.1 Materials and Solutions for Extraction of Mitochondria

1. Distilled water. 2. 0.5 M EDTA (ethylenediaminetetraacetic acid, Molecular Biology Grade, Euromedex). 3. 0.5 M EGTA (ethyleneglycoltetraacetic acid, Ultra-Pure Grade, Euromedex). 4. 1 M MgCl2 (Sigma-Aldrich). 5. DNase I (400 Kunitz units/mg protein, Sigma). 6. Percoll® (Sigma-Aldrich). 7. Stone mortar and pestle. 8. Miracloth (EMD Millipore). 9. Nylon cloth (96 μm mesh). 10. Conical flasks. 11. Fine paintbrush. 12. Dounce homogenizers. 13. Centrifuge (e.g., Avanti® J-E centrifuge, Beckman Coulter) and Rotor (e.g., JA-25.50 Beckman Coulter). 14. 50 mL centrifuge tubes and 50 mL translucent polycarbonate centrifuge tubes for the gradients. 15. Ultracentrifuge (e.g., Optima™ MAX-XP Ultracentrifuge, Beckman Coulter) and rotor (e.g., TLA-110, Beckman Coulter) (see Note 1). 16. Polycarbonate Thick Wall 5 mL tubes (Beckman Coulter). 17. Tabletop refrigerated microcentrifuge (e.g., Eppendorf 5418R with standard rotor). 18. 1.5 mL microcentrifuge tubes. 19. Balance. 20. Crushed ice.

2.2.2 Buffers for Mitochondria Isolation

1. Grinding buffer A: 0.3 M sucrose, 5 mM tetrasodium pyrophosphate, 2 mM EDTA, 10 mM KH2PO4, 1% (w/v) PVP-40, 1% (w/v) BSA, 5 mM cysteine, 20 mM sodium ascorbate, 1 mM DTT, pH 7.5 with 85% phosphoric acid (see Note 2). For the purification of mitochondria from high-water content tissues such as lettuce etiolated seedlings, use 2 × concentrated grinding buffer. About 100 mL of 1 × concentrated grinding buffer for 50 g of Arabidopsis two-weeks-old seedlings and 50 mL of 2 × concentrated grinding buffer for 12 g of oneweek-old etiolated lettuce seedlings (about 1000 seedlings).

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2. Grinding buffer B: 0.3 M sucrose, 25 mM tetrasodium pyrophosphate, 10 mM EDTA, 10 mM KH2PO4, 60 mM TES, 1 mM glycine, 1% (w/v) PVP-40, 1% (w/v) BSA, 20 mM cysteine, 50 mM sodium ascorbate, adjust to pH 8.0 with KOH. About 25 mL is required for each individual plant. 3. Washing buffer A: 0.3 M sucrose, 2 mM EGTA, 20 mM TES, 0.1% (w/v) BSA, adjust to pH 7.5 with KOH. About 400 mL is required for 50 g of Arabidopsis plants and 100 mL for 12 g etiolated lettuce seedlings (about 1000 seedlings). 4. Washing buffer B: 0.3 M sucrose, 10 mM TES, 2 mM EDTA, 10 mM KH2PO4, adjust to pH 8.0 with KOH. 2.2.3 Percoll Step Gradients for Arabidopsis Mitochondria Isolation

1. Prepare 7 mL of 5 × buffer A per gradient: 1.5 M sucrose, 5 mM EGTA, 50 mM MOPS, pH 7.2 with KOH.

2.2.4 Percoll Step Gradients for Lettuce Mitochondria Isolation

1. Prepare 600 μL of 5 × buffer L per gradient: 1.5 M sucrose, 0.5% BSA, 50 mM MOPS, pH 7.5 with KOH.

2. For each gradient, prepare 5 mL of 50% (v/v) Percoll in 1 × buffer A, 25 mL of 25% Percoll in 1 × buffer A and 5 mL of 18% Percoll in 1 × buffer A.

2. For each gradient, prepare 1 mL of 50% (v/v) Percoll in 1 × buffer L, 1.5 mL of 25% Percoll in 1 × buffer L, and 0.5 mL of 18% Percoll in 1 × buffer L.

2.3 Extraction and Purification of Nucleic Acids from Mitochondria

1. Tabletop refrigerated microcentrifuge (e.g., Eppendorf 5415R with standard rotor).

2.3.1 Common Materials and Solutions for Nucleic Acid Purification

4. TE buffer: 10 mM Tris–HCl pH 7.5, 1 mM EDTA.

2.3.2 DNA Purification Using CTAB

2. 1.5 mL microcentrifuge tubes. 3. Micropipettes. 5. NanoDrop 2000 UV–Vis spectrophotometer (ThermoScientific) for DNA quantification. 1. CTAB (Cetyltrimethylammonium bromide), Merck. 2. Chloroform/isoamylalcohol (24:1). 3. Isopropanol. 4. 70% (v/v) ethanol. 5. Dry heating block (e.g., Eppendorf Thermomixer or equivalent). 6. Fume hood.

2.3.3 DNA Purification Using QIAamp DNA Micro Kit

1. QIAamp DNA Micro kit (Qiagen).

Plant mtDNA Purification 2.3.4 RNase Treatment of Nucleic Acids

1. RNase A, 10 mg/mL (Thermo-Scientific, Massachusetts, United States).

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2. RNase T1 100 U/μL (from Aspergillus oryzae, Roche). 3. 0.2 M ammonium acetate. 4. Chloroform. 5. 70% (v/v) ethanol. 2.4 qPCR to Assess mtDNA Enrichment

1. Real-time PCR amplification instrument (e.g., Roche LightCycler 480II, Basel, Switzerland). 2. qPCR plates (e.g., LightCycler 480 Multiwell Plates 384) and sealing foils. 3. 0.5–10 μL electronic micropipette (e.g., Eppendorf Xplorer® or equivalent). 4. Eppendorf Multipette Plus pipette (or equivalent repeater pipette device) and 0.1 mL Eppendorf Combitips (1 μL minimum distribution volume). 5. Tabletop centrifuge with rotor adapted for microplates (e.g., Eppendorf 5804R and rotor A-2-DWP). 6. FastStart Universal SYBR Green Master Mix (Roche), or 2× Master Mix (Promega, Madison, Wisconsin, United States). 7. Primers: 2 μM mix of each forward and reverse qPCR primer pairs, preferentially in 0.2 mL 12-Strip tubes for convenient distribution in qPCR plates using a 12-channel electronic pipette.

3

Methods

3.1 Plant Growth Conditions 3.1.1 In Vitro Culture of Arabidopsis Plants

1. Prepare square Petri dishes (120 × 120 mm) containing autoclaved culture medium. You will need about 80 mL per dish. 2. Surface-sterilize the required amount of seeds. The equivalent to 20 μL of Arabidopsis Col-0 seeds, or 100 seeds, is enough for one Petri plate. Work in sterile conditions in a laminar flow cabinet. In a 1.5 mL tube, add 1 mL sterilizing solution to the seeds and incubate for 15 min at room temperature with agitation. Fast spin the seeds to the bottom of the tube and remove the liquid. Briefly wash twice with 70% (v/v) ethanol and once with 90% ethanol. Remove as much liquid as possible. 3. Dry the sterilized seeds in the open tube in a laminar flow cabinet. If the volume of seeds exceeds 50 μL it is advised to dry the seeds by spreading them on sterile blotting paper in a flow cabinet, because if seeds stay in ethanol for too long it will

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reduce germination efficiency. Other methods to sterilize seeds can also be used (see Note 3). 4. Distribute seeds on plates, spaced about 1 cm apart. This can be done with a sterile wooden toothpick. Seal the Petri dishes with surgical tape to retain humidity. 5. Stratify seeds by incubating plates in the dark at 4 °C for 2–3 days. Place plates upside-down to avoid condensation on the lid. 6. Grow seedlings in a growth chamber or cabinet, for approximatively 2 weeks, under a long-day photoperiod of 16 h light at 21 °C/8 h dark at 18 °C. 7. Once plants are 2 weeks old, use immediately for isolation of mitochondria (Fig. 2). 3.1.2 Arabidopsis Plants Grown in Soil

1. Sprinkle non-sterile Arabidopsis seeds onto a soil-containing pot and thoroughly water. 2. About 10 days after sowing, transplant individual plants to individual pots and grow further for about 2–3 weeks, depending on the growth rate of the accession, until a rosette size of about 6 cm diameter. 3. Cut the full rosette from the stalk and rinse with water to remove excess contaminating soil. Discard the stalky base of leaves.

3.1.3 Culture of Etiolated Lettuce Seedlings on Blotting Paper

1. Prepare ; 90 mm × 50mm Petri dishes containing one piece of sterilized T-300 blotting paper. Wet each blotting paper with 5 mL of sterile water. 2. Surface-sterilize the required amount of seeds, about 200 seeds (200 mg) per Petri dish. This should be done in a sterile laminar flow cabinet. In a 1.5 mL tube, add 1 mL of 70% ethanol to the seeds and incubate for 30 s at room temperature with agitation. Fast spin the seeds to the bottom of the tube and remove liquid. Wash with 1% NaOH for 10 min followed by three rapid washes with sterile water. Remove as much liquid as possible. 3. Disperse seeds on the wet T-300 blotting paper. Seal the Petri dishes with parafilm to retain humidity. 4. Stratify for 2 days at 4 °C to induce germination. 5. Grow seedlings in the dark (Petri dishes wrapped in aluminum foil) for a week, in a growth chamber at 21 °C. 6. Once plants are 1 week old, use immediately for isolation of mitochondria (Subheading 3.2.3) (Fig. 2).

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Fig. 2 Purification of mitochondria and qPCR quantification of mtDNA enrichment. Schematic representation of mitochondrial purification from a single Arabidopsis rosette, from Arabidopsis green seedlings, or from lettuce etiolated seedlings. The Percoll purified mitochondria are indicated by the red arrowheads. The relative enrichment of the chloroplast and mitochondrial genomes (cpDNA and mtDNA) as compared to the total plant DNA fraction was calculated by qPCR, as described. Values are normalized to the nuclear 18S rRNA gene sequence for lettuce and to the ACT1 and UBQ10 genes for Arabidopsis. Errors bars represent the SD values for biological replicates.

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3.2 Purification of Mitochondria 3.2.1 Purification of Mitochondria from Arabidopsis Seedlings

Arabidopsis mitochondria are purified mostly according to Ku¨hn et al. [10]. All steps should be performed at 4 °C with freshly prepared solutions (see Note 4). For best results, it is advised to make the gradients the day before, as it takes time. 1. Prepare gradients in 50 mL translucent centrifuge tubes by first layering the 50% Percoll solution, then slowly overlay the 25% Percoll solution with a pipette placed against the wall of the tube, 5 mm over the surface of the solution, and finish with the 18% Percoll solution by using the same technique. Store the gradient O/N at 4 °C. 2. Collect the seedlings from 12 square 120 × 120 mm Petri dishes (about 50 g for 2-weeks-old plants). A spatula with a large flat end can be convenient for this. 3. Weigh the plants and wash with cold distilled water to remove residual agar medium. 4. Grind seedlings in a cold stone mortar with 10 mL of grinding buffer A for 5 g of seedlings. Keep the residual material. 5. Filter through two pre-wet layers of miracloth into a conical flask. 6. Repeat steps 3 and 4 on the remaining material. 7. Centrifuge for 10 min at 2000 g in 50 mL tubes. 8. Transfer the supernatant into new tubes and centrifuge for 10 min at 20,000 g. 9. Gently resuspend each pellet in 1 mL of washing buffer A, with a fine paintbrush and homogenize with a Dounce homogenizer until suspension is homogeneous. 10. Dilute with washing buffer A (using half of the volume as the initial grinding buffer) and repeat steps 6 and 7. 11. Gently resuspend each pellet in a small volume of washing buffer A (1 mL for 20 g starting material) and homogenize with a small Dounce homogenizer until suspension is homogeneous. 12. Add MgCl2 to 10 mM and DNase I to 50 μg/mL. Incubate for 30 min on ice. 13. Add EDTA to 50 mM to inhibit DNase activity. 14. Layer 1.5 mL of homogenate into tubes containing 35 mL of discontinuous Percoll step gradient (see Note 5) and centrifuge for 45 min at 40,000 g without brake for the deceleration. 15. With a vacuum pump, remove the green band at the top part of the gradient (Fig. 2). 16. Collect the yellowish band at the bottom of the gradient at the 25–50% Percoll interphase (see Note 6).

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17. Divide sample into four 50 mL tubes (a maximum of 4 mL per tube) and add 36 mL of washing buffer A (see Note 7). 18. Centrifuge for 10 min at 20,000 g. Remove the supernatant with a vacuum pump and keep around 3 mL per tube (see Note 8). 19. Resuspend the loose pellets by swirling the tubes, and pool into two tubes. Fill up to 40 mL with washing buffer A. 20. Centrifuge for 10 min at 20,000 g. 21. Resuspend the pellet in 1 mL of washing buffer A, transfer into a 1.5 mL microcentrifuge tube and centrifuge for 5 min at 16,000 g. 22. Eliminate supernatant and use mitochondria pellet for DNA extraction (see Note 9). 3.2.2 Crude Partial Purification of Mitochondria from Single Arabidopsis Plants

Arabidopsis crude mitochondria purification is adapted from a protocol described by Keech et al. [11]. 1. Grind Arabidopsis rosette (about 2 g) in a cold stone mortar, first with 10 mL of grinding buffer B, then add 15 mL additional buffer and further grind until a uniform suspension is obtained. 1. Filter the homogenate through two layers of miracloth into a 50 mL centrifuge tube. 2. Centrifuge for 10 min at 1000 g to remove most of the intact chloroplasts and thylakoid membranes. 3. Transfer the supernatant into new tubes and centrifuge for 10 min at 2500 g. 4. Transfer the supernatant into new tubes and centrifuge for 20 min at 15,000 g. 5. Resuspend the pellet in 1.5 mL of washing buffer and transfer into a 2 mL microcentrifuge tube. 6. Centrifuge at 15,000 g for 20 min. 7. Remove the supernatant and use the crude mitochondrial pellet for DNA extraction.

3.2.3 Purification of Mitochondria from Etiolated Lettuce Seedlings

Lettuce mitochondria are purified mostly according to Fertet et al. [12]. 1. Prepare gradients in 5 mL translucent tubes by first layering the 50% Percoll solution, then slowly overlay the 25% Percoll solution with a micropipette placed against the wall of the tube just over the surface of the solution, and finish with the 18% Percoll solution by using the same technique. Store the gradient at 4 ° C until use.

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2. Collect approximately 1000 etiolated seedlings and weigh them (approximately 12 g). 3. Grind the seedlings in a cold mortar using 10 mL of 2 × concentrated grinding buffer A for 200 seedlings. 4. Filter through one layer of nylon cloth (96 μm mesh) and one layer of miracloth (pre-wet with grinding medium) into a conical flask. Keep the residual material. 5. Repeat steps 2 and 3 on the residual material. 6. Centrifuge the filtrate for 10 min at 3500 g. 7. Transfer the supernatant into new tubes and centrifuge again for 5 min at 3500 g. 8. Transfer the supernatant into new tubes and centrifuge for 5 min at 6000 g. 9. Transfer the supernatant into new tubes and centrifuge for 20 min at 17,000 g. 10. Gently resuspend each pellet in a small volume of washing buffer A (200 μL per 5 g starting material) and homogenize with a mini Dounce until suspension is homogeneous. 11. Layer 200 μL on top of 3.0 mL Percoll step gradients (see Note 10). 12. Centrifuge for 25 min at 25,000 g without brake for deceleration. 13. Remove the band in the top part of the gradients with a micropipette (Fig. 2). 14. Collect the yellowish bands at the 25–50% Percoll interface. 15. Add 30 mL of washing buffer A. 16. Centrifuge for 15 min at 18,000 g. 17. Repeat the washing step. 18. Resuspend the pellet in 200 μL of washing buffer A. 19. Transfer into a microcentrifuge tube, add MgCl2 and 100 μg of DNase I. 20. Incubate for 20 min at 25 °C. 21. Add 1 mL of washing buffer A containing 10 mM EDTA and 10 mM EGTA. 22. Centrifuge 15 min at 16,000 g to sediment mitochondria, and remove the supernatant. 23. Use the pellet immediately for DNA extraction or freeze in liquid nitrogen and store at -80 °C.

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3.3 Extraction and Purification of Nucleic Acids from Mitochondria

1. Resuspend the mitochondrial pellet in preheated 300 μL CTAB lysis buffer and incubate for 45 min at 65 °C.

3.3.1 Mitochondrial DNA Purification Using CTAB

3. Centrifuge for 10 min at 12,000 g at room temperature.

2. In a fume hood, add 1 volume of chloroform/isoamylalcohol (24:1) and mix. 4. Transfer the upper phase into a new microcentrifuge tube, add 1 volume of isopropanol, mix gently and incubate for 10 min on ice. 5. Centrifuge for 10 min at 16,000 g. 6. Discard the supernatant and add 1 mL of 70% (v/v) ethanol. 7. Centrifuge for 2 min at 16,000 g. 8. Discard the supernatant, dry the pellet at room temperature and dissolve it in 20–50 μL of TE buffer. 9. Quantify the DNA concentration of samples on a spectrophotometer (e.g., NanoDrop spectrophotometer). Precise DNA quantification for the preparation of NGS libraries is done with a Qubit fluorometer. DNA extracted by the CTAB method is often contaminated with residual RNA which interferes with spectrophotometric quantification of DNA. Alternatively, analyze samples by electrophoresis of an aliquot on a 0.8% agarose gel, and adjust samples to comparable concentrations according to the intensity of the ethidium bromide-stained high molecular weight DNA band. If RNAs need to be eliminated, perform an RNase treatment as described in Subheading 3.3.3. The purification of DNA using CTAB from mitochondria of 7 day old lettuce seedlings yields about 40 ng of enriched mtDNA from 200 seedlings.

3.3.2 Mitochondrial DNA Purification Using QIAamp DNA Micro Kit (Qiagen)

For many projects, the final goal of mtDNA purification is to do whole genome sequencing for de novo assembly of the mtDNA of a species or variety that has not been studied before, or to identify sequence variants or mtDNA rearrangements. For purification of high-quality mtDNA, as required for the preparation of NGS libraries and sequencing, it is advised to use a commercial kit or an equivalent resin-based DNA purification protocol. We obtained good results with the QIAamp DNA Micro kit. For this method, resuspend the mitochondrial pellet in lysis buffer and follow the procedure of the kit. With the use of this kit, about 300 ng of enriched mtDNA can be obtained when starting from 1000 lettuce seedlings or from 50 g of two-week old Arabidopsis seedlings. When performing crude mitochondrial purification, about 20 ng of DNA was recovered from 2 g of sample (whole Arabidopsis rosette).

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For the construction of libraries for illumina whole mtDNA sequence, we routinely use the Nextera™ DNA Flex Library Preparation Kit (Illumina®), starting from 50 ng of DNA, as precisely determined by Qubit™ Fluorometric Quantification. The average size of the libraries obtained with this kit is of about 500–600 bp. 3.3.3 RNase Treatment of Nucleic Acids

If needed, after CTAB purification of the mtDNA, the contaminant RNA can be removed by RNase treatment. 1. Incubate the nucleic acid solution for 1 h at 37 °C in the presence of 1 U/μL RNase T1 and 0.1 μg/μL RNase A. 2. Add ammonium acetate to 0.2 M and add 1 volume of phenol/ chloroform (v/v). 3. Mix by agitation for 5 min. 4. Centrifuge for 10 min at 15,000 g and transfer the aqueous phase into a new tube. 5. Add 1 volume of chloroform. 6. Mix for 30 s. 7. Centrifuge 10 min at 15,000 g and transfer the aqueous phase into a new tube. 8. Add 2.5 volumes of ethanol and incubate for 30 min at -20 ° C. 9. Centrifuge for 10 min at 12,000 g. 10. Discard the supernatant and add 1 mL of 70% (v/v) ethanol. 11. Centrifuge 2 min at 16,000 g. 12. Eliminate the supernatant, dry the nucleic acid pellet at room temperature, and resuspend in a small volume (20–50 μL) of TE.

3.4 Analyses of mtDNA Enrichment by qPCR

To evaluate mtDNA enrichment, the relative levels of nuclear, plastidial, and mitochondrial DNA can be determined by qPCR. The procedure outlined here has also been described in a recent paper from this collection [13]. 1. Design primers for the amplification of unique target sequences from the nuclear, plastidial, and mitochondrial genomes (see Notes 11 and 12). It is important to select sequence regions where DNA polymorphisms are rarely found, to avoid possible mismatches at the qPCR primers binding sites that will affect the amplification efficiency. The specificity of the primers should be checked by BLAST search (http://www.ncbi.nlm. nih.gov/blast) to ensure that the targeted sequences are single copy in the genomes. The primers we routinely use are given in Table 1.

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Table 1 Table of primers used for quantification of mtDNA enrichment

Gene name

Primer sequences

Genome specificity

Primers used to test mt DNA enrichment in Arabidopsis ACT1

CATCTTGGCCTCCCTCAGTA and GAGTAAACAAGTGATGGGACTGTG

Nuclear

UBQ10

TCACCGGAAAGACCATCACT and CGGTGGGATACCCTCTTTG

Nuclear

clpP

TCGCACTATATGTCAACCCAAG and CTATTGGCGTTCCAAAAGTACC

Chloroplastic

ndhH

TCCGGATAAACCCCAATTTA and AGAACGGGTTGAAGGAGTTG

Chloroplastic

COX2

AATAAACGTGATTGACCCAATTCT and TCCGATGAGCAGTCACTCAC

Mitochondrial

rrn18S

CCTTGAGCTAGGAGCCTCTTT and CATGCAAGTCGAACGTTGTT

Mitochondrial

Primers used to test mt DNA enrichment in Lactuca sativa rrn18S

ACTCCGCTGGCACCTTATGAG and GTGGTGCCCTTCCGTCAATTC

Nuclear

rbcL

CAGTTCGGTGGAGGAACTTTAG and GCAAGATCGCGTCCCTATTAC

Chloroplastic

mt encoded rrn18S

CGAGTGCGCGATCATGACAAG and CGCCCAGTCATTCCGAAGAAC

Mitochondrial

2. Prepare qPCR plates. For the LightCycler 480II, use 384-well white qPCR plates. Each reaction is performed in triplicate, in a total reaction volume of 6 μL. During setup, plates are kept cold on ice, to avoid evaporation (see Note 13). For ease of distribution, use multichannel pipettes and prepare all mixes in strips of 0.2 mL tubes. 3. Distribute 2 μL of each primer mix (2 μM of forward and reverse primers) with a Multipette. For instance, in a 384-well plate you can distribute up to eight different primer pairs vertically (columns 1–3, 4–6, 7–9, 10–12, 13–15, 16–18, 19–21, 22–24 of the plate) for the analysis of up to 16 samples, distributed horizontally (rows A to P). One of the samples should be a negative control of amplification containing no added DNA. Briefly spin plates before continuing. 4. For each reaction, prepare a mix of 3 μL of SYBR 2 × Master Mix and 1 μL of diluted DNA sample (or water in the negative control). For a total of 24 reactions, 10 ng of total plant DNA is sufficient, and much less from purified mtDNA (see Note 14).

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5. Distribute 4 μL of each mix to wells with a Multipette, or with a multichannel automatic pipette in the mode of distribution, if found more convenient. Cover plate with sealing foil, and centrifuge briefly. 6. Run PCR program. The parameters we routinely use are: 7 min at 95 °C, to activate the hot-start enzyme and denature the DNA, followed by 40 cycles of 10 s at 95 °C, 15 s at 58 °C, and 15 s at 72 °C. At the end of the PCR cycles, determine the Tm values by melting curve analysis, following the default program of the LightCycler 480II. A specific PCR reaction will produce an amplicon with just a single Tm value. 7. For each sample and primer pair, from the technical triplicates calculate the average Cp value and standard deviation (SD). Ideally, the SD should not exceed 0.2 Cp units. 8. Calculate relative concentrations using the ΔΔCp method. Briefly, for each amplified sequence, the relative concentration in a sample (as compared to the reference sample, e.g., WT) is given by ε-ΔCp, where ΔCp = (average Cp of sample)-(average Cp of WT) and ε is the amplification efficiency. In most cases where precision is not essential, it can be assumed that amplification efficiency is 100% (ε = 2). The estimation of ε can be done using LineRegPCR (http://LinRegPCR.nl). 9. Normalize the relative concentrations to the nuclear genome, by dividing by a correction factor that is the average of the concentration values obtained for the nuclear sequences. Figure 2 represents the relative enrichment in organellar genomes in the DNA extracted from purified mitochondria of Arabidopsis or lettuce, as compared to total plant DNA. Table 2 shows the values of coverage obtained from Illumina sequencing.

Table 2 Coverage of the organellar genomes on Illumina reads from DNA extracted from crude and Percoll gradient purified mitochondria Arabidopsis 4-week plant Total plant DNA (%)

Arabidopsis 4-week plant Partially purified mtDNA (%)

Arabidopsis 2-week seedlings Purified mtDNA (%)

Lettuce etiolated seedling Purified mtDNA (%)

cpDNA 21

3

41

7

mtDNA 2

13

40

53

Crude purification from a single Arabidopsis plant results in a sixfold increased coverage of the mtDNA as compared to total plant DNA (13% versus 2%). Percoll gradient purification of Arabidopsis mitochondria results in a 20-fold increase in mtDNA, but also a significant increase in chloroplastic DNA, because immature proplasts co-purify with mitochondria. The mtDNA purified from lettuce etiolated seedlings is less contaminated with chloroplastic DNA

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3.5 Analyses of mtDNA Enrichment from Illumina Whole Genome Sequencing Data

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While qPCR gives an assessment of the relative mtDNA enrichment as compared to the nuclear or chloroplast genomes, absolute values can be calculated from the total number of NGS reads mapping to each one of the three plant genomes, starting from either total plant DNA or from purified mtDNA. This requires that the sequences of the corresponding reference genomes are known. The mapping of Illumina reads to a reference genome is beyond the scope of this chapter. However, we routinely use the software “bowtie” (http://bowtie-bio.sourceforge.net/index.shtml), which is implemented in MacVector Assembler (https://macvector.com/ Assembler/assembler.html). Because most plant mitogenomes contain sequences of chloroplast origin, we first map the reads to the cpDNA, which has a much higher ploidy than the mtDNA. The reads that do not match the cpDNA are then mapped to the mtDNA. This results in a slight undervaluation of the mtDNA coverage, depending on the cpDNA contamination. Table 2 shows the values of mtDNA and cpDNA coverage obtained from Illumina sequencing data, starting from about four million total reads in each case. This example shows that the mtDNA sequences account for just 2% of the total Arabidopsis DNA, but in a Percollpurified mtDNA fraction about 40% of the DNA is mitochondrial.

Notes 1. A high-speed centrifuge and rotor for 50 mL tubes is needed when starting with relatively large amounts of material, requiring the centrifugation of 35 mL Percoll gradients. For small tissue amounts, purification can be carried out on 3 mL Percoll gradients prepared in 5 mL centrifuge tubes spun on a benchtop ultracentrifuge, as described for lettuce etiolated seedlings. 2. The day before the mitochondrial isolation, dissolve sucrose, tetrasodium pyrophosphate, and EDTA, adjust to pH 7.5 and then add BSA and PVP. Store overnight at 4 °C and add cysteine and sodium ascorbate before use. 3. Vapor-phase seed sterilization is convenient for the simultaneous surface-sterilization of many seed samples. Briefly, place open tubes in a rack inside a vacuum desiccator in a fume hood. Place a 250 mL beaker containing 50 mL of bleach (1% (w/v) sodium hypochlorite) inside the desiccator. Using a glass pipette, add 3 mL of concentrated HCl (37%) into the bleach and rapidly close the desiccator. Leave for about 4 h up to overnight. 4. The whole purification process must be done at 4 °C. If possible, the grinding is performed in a cold room. Tubes and rotors must be precooled before use. Grind in several times, using 5 g of tissue at a time.

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5. For each gradient, use a maximum amount of extract corresponding to 30 g starting material. 6. When collecting from the Percoll gradient, it is advised to leave the bottom 2–3 mL of the gradient. 7. If Percoll is present in too high concentration, then mitochondria will not sediment during centrifugation, so do not exceed 1/10 volume of mitochondria recovered from the gradient per total volume. 8. Be careful because the pellet will be loose. 9. If necessary, mitochondrial pellets can be frozen in liquid nitrogen and stored at -80 °C before extracting nucleic acids. 10. For each gradient, use a maximum starting amount corresponding to 5 g starting material. 11. It should be also taken into account that there are sequences of mitochondrial origin in the nuclear genome (NUMTs, for “nuclear mitochondrial DNA“). Thus, if the nuclear genome sequence is known, it is important to select regions which are only found in the mtDNA. When studying the mtDNA of species whose nuclear genomes have not yet been sequenced, the importance of NUMTs cannot be ignored. 12. Primer pairs should have equivalent melting temperatures (Tm), of about 60–65 °C, and a G + C composition of 45–55%. The size of the amplicon should be around 70–120 bp. 13. If the preparation of the qPCR plate takes too long, the evaporation of samples can change the concentrations of the reaction mixes. It is important to adopt rapid and efficient strategies of preparing the plates. Evaporation can be kept minimal by keeping plates on ice, or preparing in a cold room. A convenient method to keep qPCR plates cold and dry is to cover the ice with a rigid and flat aluminum plate, and place the qPCR plate over the cold aluminum plate. 14. It is very important to thoroughly mix the solutions, because the SYBR 2 × Master Mix is viscous. If not, there is the risk of distributing different amounts of DNA sample in the replicates. Mixing can be done by vortexing for 30 s, if the mixes are prepared in individual 1.5 mL tubes, or by repeated pipetting up and down if working with 0.2 mL tube strips.

Acknowledgments This work was supported by the LABEX MitoCross [ANR-11LABX-0057_MITOCROSS] and benefited from a funding managed by the French National Research Agency as part of the “Investments for the future” program.

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References 1. Moustafa IM, Uchida A, Wang Y, Yennawar N, Cameron CE (2015) Structural models of mammalian mitochondrial transcription factor B2. Biochim Biophys Acta 184:987–1002. https://doi.org/10.1016/j.bbagrm.2015. 05.010 2. Sloan DB, Alverson AJ, Chuckalovcak JP, Wu M, McCauley DE, Palmer JD, Taylor DR (2012) Rapid evolution of enormous, multichromosomal genomes in flowering plant mitochondria with exceptionally high mutation rates. PLoS Biol 10:e1001241. https:// doi.org/10.1371/journal.pbio.1001241 3. Wynn EL, Christensen AC (2019) Repeats of unusual size in plant mitochondrial genomes: identification, incidence and evolution. G3 (Bethesda) 9:549–559. https://doi.org/10. 1534/g3.118.200948 4. Preuten T, Cincu E, Fuchs J, Zoschke R, Liere K, Bo¨rner T (2010) Fewer genes than organelles: extremely low and variable gene copy numbers in mitochondria of somatic plant cells. Plant J 64:948–959. https://doi. org/10.1111/j.1365-313X.2010.04389.x 5. Takanashi H, Ohnishi T, Mogi, Okamoto T, Arimura S, Tsutsumi N (2010) Studies of mitochondrial morphology and DNA amount in the rice egg cell. Curr Genet 56:33–41. https://doi.org/10.1007/s00294-0090277-3 6. Sodmergen, Zhang Q, Zhang Y, Sakamoto W, Kuroiwa T (2002) Reduction in amounts of mitochondrial DNA in the sperm cells as a mechanism for maternal inheritance in Hordeum vulgare. Planta 216:235–244. https:// doi.org/10.1007/s00425-002-0853-y 7. Hazkani-Covo E, Zeller RM, Martin W (2010) Molecular poltergeists: mitochondrial DNA copies (numts) in sequenced nuclear genomes.

PLoS Genet 6:e1000834. https://doi.org/10. 1371/journal.pgen.1000834 8. Stupar RM, Lilly JW, Town CD, Cheng Z, Kaul S, Buell CR, Jiang J (2001) Complex mtDNA constitutes an approximate 620-kb insertion on Arabidopsis thaliana chromosome 2: implication of potential sequencing errors caused by large-unit repeats. Proc Natl Acad Sci U S A 98:5099–5103. https://doi.org/ 10.1073/pnas.091110398 9. Galbraith DW, Harkins KR, Knapp S (1991) Systemic endopolyploidy in Arabidopsis thaliana. Plant Physiol 96:985–989. https://doi. org/10.1104/pp.96.3.985 10. Ku¨hn K, Obata T, Feher K, Bock FAR, Meyer EH (2015) Complete mitochondrial complex I deficiency induces an up-regulation of respiratory fluxes that is abolished by traces of functional complex I. Plant Physiol 168:1537– 1549. https://doi.org/10.1104/pp.15. 00589 11. Keech O, Dizengremel P, Gardestrom P (2005) Preparation of leaf mitochondria from Arabidopsis thaliana. Physiol Plant 124:403– 409. https://doi.org/10.1111/j.1399-3054. 2005.00521.x 12. Fertet A, Graindorge S, Koechler S, de Boer GJ, Guilloteau-Fonteny E, Gualberto JM (2021) Sequence of the mitochondrial genome of Lactuca virosa suggests an unexpected role in Lactuca sativa’s evolution. Front Plant Sci 12:697136. https://doi.org/10.3389/fpls. 2021.697136 13. Schatz-Daas D, Fertet A, Lotfi F, Gualberto JM (2022) Assessment of mitochondrial DNA copy number, stability and repair in Arabidopsis. Methods Mol Biol 2363:301–319. https:// doi.org/10.1007/978-1-0716-1653-6_20

Part II Visualising mtDNA

Chapter 6 Visualize the Distribution and Dynamics of Mitochondrial DNA (mtDNA) Nucleoids with Multiple Labeling Strategies Xiangjun Di, Jinshan Qin, Yujie Sun, and Qian Peter Su Abstract Mitochondrial DNA (mtDNA) encodes a variety of rRNAs, tRNAs, and respiratory chain complex proteins. The integrity of mtDNA supports the mitochondrial functions and plays an essential role in numerous physiological and pathological processes. Mutations in mtDNA cause metabolic diseases and aging. The mtDNA within the human cells are packaged into hundreds of nucleoids within the mitochondrial matrix. Knowledge of how the nucleoids are dynamically distributed and organized within mitochondria is key to understanding mtDNA structure and functions. Therefore, visualizing the distribution and dynamics of mtDNA within mitochondria is a powerful approach to gain insights into the regulation of mtDNA replication and transcription. In this chapter, we describe the methods of observing mtDNA and its replication with fluorescence microscopy in both fixed and live cells using different labeling strategies. Key words Mitochondrial DNA (mtDNA), TFAM, POLG2, BrdU, EdU, PdG

1

Introduction Hundreds of copies of mitochondrial DNA (mtDNA) are organized into nucleoids in mitochondria within the human cell cytoplasm. Dynamic replication, segregation, and distribution are crucial to mtDNA maintenance and mitochondrial functions [1– 3]. Dysfunction of mtDNA integrity has been found to be related to a wide range of diseases, such as neurodegenerative and senescence-linked disorders [2, 4]. Using fluorescence microscopy, Lewis et al. found that mtDNA replication often occurs at the endoplasmic reticulum (ER)-mitochondria contact sites where mitochondrial division takes place [5]. With the recently developed lattice light-sheet microscopy, McArthur et al. demonstrated that the Bcl-2 homologous antagonist killer and Bcl-2-associated X protein (BAK/BAX) macropores facilitate mtDNA efflux during

Xiangjun Di and Jinshan Qin contributed equally to this work. Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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the apoptosis process [6]. More recently, using live-cell super-resolution microscopy, Qin et al. found that the mtDNA nucleoids in mitochondria could be actively transported by kinesin family member 5B (KIF5B)-driven mitochondrial dynamic tubulation activities [7]. Therefore, directly observing mtDNA replication and its dynamics via advanced imaging techniques has facilitated the mechanistic understanding of mtDNA distribution within cells. The ideal approaches for labeling and visualizing mtDNA and its replication, in both live and fixed cells, are expected to satisfy the following properties [8]: • Specific labeling toward mtDNA rather than nuclear DNA (~16 kilos vs 3 billion base pairs). • Limited effects on the normal dynamics and functions of mtDNA in live cells. • Perfect photostable performance for long-time observation and tracking. • Flexibility in the spectral characteristics for multi-color imaging. • Preferred targeting to replicating mtDNA selectively. • Good compatibility with multiple super-resolution microscopy systems to achieve high spatiotemporal resolution. • Labeling sequence variants of mtDNA selectively, including single-nucleotide variants, to help understand the pathophysiology of mtDNA heteroplasmy. These properties are desirable but challenging. Here, we summarize the protocols for tracking total mtDNA and its replication with fluorescent protein-tagged mitochondrial transcription factor A (TFAM) and DNA polymerase gamma 2 (POLG2), respectively, in live cells. We also summarize the methods of mapping total mtDNA and its replication in fixed cells by different labeling strategies, including immunofluorescence staining assays toward TFAM/DNA, DNA POLγ, and BrdU (thymidine analogue, 5-bromo-2′-deoxyuridine), as well as Click-iT reactions for labeling of EdU (thymidine analogue, 5-ethynyl-deoxyuridine) and PdG (guanine analogue, 6-O-propynyl-2′-deoxyguanosine), which is a selective mitochondrial DNA staining strategy.

2

Materials Prepare all solutions and buffers using analytical grade reagents and dissolve in ddH2O, then filter with 0.2 μm syringe filters to remove impurities (which may auto-fluoresce during fluorescence imaging). Prepare and store all reagents, solutions, and buffers at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing of waste materials. Do not add sodium azide (NaN3) to the solutions.

Distribution and Dynamics of mtDNA

2.1 Observation of Total mtDNA 2.1.1 Map Total mtDNA Distribution in Fixed Cell

81

1. ddH2O: MilliQ, Millipore, 18.2 MΩ cm at 25 °C. 2. Culture medium: DMEM (high glucose) supplemented with 10% FBS, 1% penicillin, 1% streptomycin, and GlutaMAX-ITM. 3. Phosphate-buffered saline (PBS): pH 7.4, 135 mM NaCl, 4.7 mM KCl, 10 mM Na2HPO4, 2 mM NaH2PO4. 4. PHEM buffer: 60 mM Pipes pH 6.9, 25 mM Hepes, 10 mM EGTA, and 2 mM MgCl2. 5. 10% bovine serum albumin (BSA) (w/v) in PBS. 6. Blocking buffer: 0.5% Triton X-100 (v/v) and 1.0% BSA (w/v) in PHEM buffer. 7. Primary antibody: anti-TFAM, anti-DNA antibody. Store at 20 °C or follow the manufacturer’s manual before using. 8. Secondary antibody: dye-conjugated IgG which fits the wavelength of the microscopy channel. Store at -20 °C. 9. Cell culture dish: Fluorodish 35 mm with No.1 (thickness = 170 μm) coverglass. 10. 6% paraformaldehyde (PFA) in PHEM buffer.

2.1.2 Track Total mtDNA Dynamics in Live Cell

1. ddH2O, culture medium, PBS, and cell culture dish (same as in Subheading 2.1.1), 2. Plasmid: (1) mtDNA nucleoids marker: TFAM-GFP, TFAMmCherry; (2) mitochondrial marker: TOM20-GFP, Mito-YFP, Mito-BFP. 3. Opti-MEM medium. 4. Lipofectamine 2000 reagent. 5. Fibronectin. 6. Quant-iT™ PicoGreen™ dsDNA Reagent.

2.2 Observation of mtDNA Replication 2.2.1 Map mtDNA Replication in Fixed Cell

1. ddH2O, culture medium, PBS, PHEM buffer, BSA, blocking buffer, cell culture dish, 6% PFA, and secondary antibody (same as in Subheading 2.1.1), 2. 5-Bromo-2′-deoxyuridine (BrdU) powder. Store at -20 °C before dissolving in PBS or ddH2O. 3. 5-Ethynyl-dU (EdU) powder. Store at -20 °C before dissolving in dimethyl sulfoxide (DMSO). 4. 6-O-propynyl-dG (PdG) powder. Store at -20 °C before dissolving in DMSO. 5. Primary antibody: anti-BrdU, anti-DNA POLγ antibody. Store at -20 °C or follow the manufacturer’s manual before using. 6. 2 M HCl in PBS or ddH2O. 7. Tween 20.

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8. 0.5% Triton X-100 in PBS. 9. Click-iT™ Cell Reaction Buffer Kit: Click-iT® cell reaction buffer (Component A), copper(II) sulfate (CuSO4, Component B), Click-iT® cell buffer additive (Component C). Store at 4 °C. 10. CuAAC Reaction Ligand Test Kit (THPTA): CuSO4, THPTA stock solution, Na-Ascorbate stock solution. Store at 4 °C. 11. Dye-azide dissolved in DMSO. Store at -20 °C. 2.2.2 Track mtDNA Replication in Live Cell

1. ddH2O, culture medium, PBS, Opti-MEM medium, Lipofectamine 2000 reagent, fibronectin, and cell culture dish (same as in Subheading 2.1.1), 2. Plasmid: (1) mtDNA replication marker: POLG2-GFP; (2) mitochondrial marker: TOM20-GFP, Mito-YFP, Mito-BFP.

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Methods

3.1 Observation of Total mtDNA

1. Culture cells in a Fluorodish with culture medium at 37 °C in the presence of 5% CO2.

3.1.1 Map Total mtDNA Distribution in Fixed Cell

2. The next day, check the cell coverage and wash the cells three times in PBS. 3. Fix the cells with 6% PFA in PHEM buffer for 15 min at room temperature. Then wash the cells 3 times with PHEM buffer. 4. Incubate the cells for 30 min at room temperature with the blocking buffer. 5. Incubate the cells with primary antibody anti-TFAM or antiDNA antibody (with the dilution ratio recommended by the manufacturer) in blocking buffer overnight at 4 °C. 6. Wash the cells with PHEM buffer three times for 5 min each wash. 7. Incubate the cells with dye-labeled secondary antibody (1:50– 1:200) for 1 h at room temperature (see Note 1). 8. Wash with PHEM buffer three times for 5 min each wash. 9. Post-fix the samples with 6% PFA in PHEM buffer for 10 min at room temperature. 10. Image mtDNA using a fluorescence microscope (Fig. 1a).

3.1.2 Track Total mtDNA Dynamics in Live Cells Track Total mtDNA in Live Cells by Transfecting with Plasmids

1. Incubate the dish with 5 μg/cm2 fibronectin. 2. One day before transfection, plate 0.5–2 × 105 cells in 500 μL growth medium without antibiotics so that cells will be 70–90% confluent at the time of transfection.

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Fig. 1 Observation of total mtDNA. (a) Confocal image of cells immuno-stained with anti-mtDNA antibody. (b) Live-cell confocal image of a cell transfected with mito-YFP (red) and TFAM-mCherry (green) plasmid. (c) Livecell confocal image of a cell treated with PicoGreen

3. Dilute 2 μg of plasmid DNA (TFAM-GFP, TFAM-mCherry, etc.) in 50 μL Opti-MEM medium. 4. Mix Lipofectamine 2000 gently before use, then dilute 5 μL Lipofectamine 2000 reagent in 50 μL Opti-MEM medium. Incubate for 5 min at room temperature (see Note 2). 5. Combine the diluted DNA plasmid and diluted Lipofectamine 2000. Mix gently and incubate for 20 min at room temperature (see Note 3). 6. Add the mixture to the dish. Mix gently by rocking the dish back and forth. 7. Incubate cells at 37 °C in a CO2 incubator for 16–36 h before imaging. The medium may be changed after 4–6 h. 8. Image the cells with a fluorescence microscope (Fig. 1b). Track Total mtDNA in Live Cells by Staining with PicoGreen dsDNA Reagent

1. Dilute PicoGreen DNA with culture medium (1:1000). 2. Incubate the cells with PicoGreen dilution at 37 °C in a CO2 incubator for 15 min. 3. Wash the cells three times in PBS. 4. Image the cells with a fluorescence microscope (Fig. 1c).

3.2 Observation of mtDNA Replication 3.2.1 Map mtDNA Replication in Fixed Cell Immunostaining Toward POLγ

1. Repeat steps 1–10 in Subheading 3.1.1 with anti-DNA POLγ antibody.

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Incorporation of BrdU, EdU, and PdG

1. Prepare a 10 mM stock solution of BrdU by dissolving 3 mg BrdU in 1 mL ddH2O. Prepare a 10 mM stock solution of EdU by dissolving 2.52 mg EdU in 1 mL DMSO. Prepare a 10 mM stock solution of PdG by dissolving 3 mg PdG in 1 mL DMSO. 2. Culture cells in culture medium at 37 °C in the presence of 5% CO2. 3. Dilute the 10 mM BrdU/EdU/PdG stock solution in cell culture medium to make a 10–100 μM BrdU/EdU/PdG labeling solution (see Note 4). 4. The following day, remove the culture medium from the cells and replace it with the labeling solution (see Note 5). 5. Incubate the cells in the BrdU/EdU/PdG labeling solution for 1–24 h at 37 °C in the presence of 5% CO2 (see Note 6). 6. Remove the BrdU/EdU/PdG labeling solution from the cells and wash twice in PBS for 5 s per wash. Wash three more times with PBS for 2 min each time. 7. Perform sequential labeling experiments by completely removing media with the first thymidine analog, rinsing twice in fresh media, and replacing with media containing the second thymidine analog.

BrdU Labeling

1. Fix cells with 6% PFA in PBS pH 7.4 for 15 min at room temperature. 2. Wash three more times with PBS for 2 min each. 3. Incubate cells in 2 M HCl in 0.1% PBS-Tween for 30 min at room temperature to denature the double-stranded DNA and expose the BrdU epitope (see Note 7). 4. Repeat steps 4–9 in Subheading 3.1.1 with anti-BrdU antibody. 5. Image mtDNA using a fluorescence microscope (Fig. 2a).

EdU Labeling

1. Fix cells with 6% PFA in PBS pH 7.4 for 15 min at room temperature. 2. Remove the fixative and wash the cells twice with 1 mL 3% BSA in PBS. 3. Remove the washing solution. Incubate the cells with 1 mL 0.5% Triton X-100 in PBS (permeabilization buffer) for 20 min. 4. Prepare 1× Click-iT EdU buffer additive (see Note 8). 5. Prepare a Click® reaction cocktail: mix 430 μL 1× Click-iT® reaction buffer, 20 μL CuSO4, 1.2 μL dye-azide, and 50 μL reaction buffer (see Note 9).

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Fig. 2 Observation of mtDNA replication. (a) Total Internal Reflection Fluorescence (TIRF) image of a cell labeled with EdU (magenta, image contrast has been enhanced to see the mtDNA replication, while the nucleus DNA is overexposed) and immuno-stained with anti-TFAM (green) antibody. (b) TIRF image of cells labeled with BrdU and immuno-stained with anti-BrdU (magenta) and anti-TFAM (green) antibodies

6. Remove the permeabilization buffer and wash twice in 3% BSA in PBS. Remove the washing solution. 7. Add 500 μL of Click-iT® reaction cocktail to each dish. Incubate the dish for 30 min at room temperature, avoiding light. 8. Remove the reaction cocktail, then wash the cells twice with 3% BSA in PBS. 9. Post-fix the samples with 6% PFA in PBS for 10 min at room temperature. 10. Image mtDNA using a fluorescence microscope (Fig. 2b). PdG Labeling

1. Fix cells with 6% PFA in PBS pH 7.4 for 15 min at room temperature. 2. Remove the fixative and wash the cells twice with 1 mL of 3% BSA in PBS. 3. Remove the washing solution. Incubate the cells with 1 mL 0.5% Triton X-100 in PBS (permeabilization buffer) for 20 min.

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4. Prepare CuSO4:THPTA-Premix: mix 10 μL 100 mM CuSO4 and 20 μL 250 mM THPTA stock solution by vortexing and spin down briefly. 30 μL of CuSO4:THPTA-Premix is sufficient for the preparation of 500 μL of Click reaction cocktail (see Note 10). 5. Prepare Click reaction cocktail: add 1 μL 10 mM azide or alkyne detection reagent solution to 419 μL reaction buffer, vortex and spin down briefly. Add 30 μL CuSO4:THPTAPremix, vortex, and spin down briefly. Add 50 μL Na-Ascorbate, vortex, and spin down briefly (see Note 11). 6. Remove the permeabilization buffer and wash twice in 3% BSA in PBS. Remove the washing solution. 7. Add 500 μL Click reaction cocktail to the Fluorodish and incubate for 30–60 min at room temperature, protected from light. 8. Remove the reaction cocktail, then wash the cells twice with 3% BSA in PBS. 9. Post-fix the samples with 6% PFA in PBS for 10 min at room temperature. 10. Image mtDNA using a fluorescent microscope. Dual BrdU and EdU/PdG Labeling

1. Fix cells with 500 μL 6% PFA for 10 min at room temperature and rinse twice in PBS. 2. Add 1 mL 0.5% Triton X-100 in PBS for 10 min at room temperature. 3. Label EdU/PdG first, following the protocols described in Subheadings 3.2.1.4 and 3.2.1.5, as HCl treatment does not compromise the EdU/PdG signal. 4. Add 2 M HCl to the dish for 30 min at 37 °C to recover the BrdU epitope. 5. Wash the cells three times in PBS. 6. Label the BrdU following the protocol described in Subheading 3.2.1.3. 7. Post-fix the samples with 6% PFA in PBS for 10 min at room temperature. 8. Image mtDNA using a fluorescent microscope.

3.2.2 Observation of mtDNA Replication in Live Cells

1. Repeat steps 1–8 in Subheading 3.1.2.1 with POLG2-GFP plasmid.

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Notes 1. Avoid light for the secondary antibody. 2. Dilute Lipofectamine 2000 freshly and use it within 25 min. 3. The DNA plasmid and Lipofectamine 2000 complexes are stable for 6 h at room temperature. 4. For BrdU/EdU labeling, the exact concentration should be optimized for your experiment. It is hard to see mtDNA staining with 10 μM BrdU/EdU, as they prefer to incorporate in nucleus DNA first. A final labeling solution concentration of 100 μM is recommended here. 5. Do not wash cells just before incubation with BrdU. This will slow the growth of cells during the incorporation phase of the procedure. 6. BrdU incubation time depends on how rapidly cells divide. Primary cells may need up to 24 h, while rapidly proliferating cell lines may only need 1 h. The exact duration should be optimized to achieve the optimal signal-to-noise ratio. 7. The exact HCl concentration and incubation time should be optimized for your experiment. If incubating at 37 °C, the incubation time should be shorter. 8. Prepare the Click-iT EdU buffer additive solution freshly and use it on the same day. 9. The ingredients are introduced in the order presented; otherwise, the reaction will not proceed optimally. Use the ClickiT® reaction cocktail within 15 min of preparation. 10. Prepare CuSO4:THPTA-Premix freshly for each experiment. Warm up all the solutions to room temperature before use in reactions. 11. Prepare the Click-iT reaction cocktail freshly for each experiment and use it within 15 min of preparation.

Acknowledgments This work was supported by the funding from the National Key R&D Program of China (No.2017YFA0505300) and the National Science Foundation of China (21825401) to Y.S.; grants from the Australia National Health and Medical Council (NHMRC, APP1177374) and the Australia National Heart Foundation (NHF, 102592) to Q.P.S.; and the funding from China Scholarship Council (No. 201706170028) to X.D.

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References 1. Garrido N et al (2003) Composition and dynamics of human mitochondrial nucleoids. Mol Biol Cell 14(4):1583–1596 2. Friedman JR, Nunnari J (2014) Mitochondrial form and function. Nature 505(7483):335–343 3. Wang C et al (2015) Dynamic tubulation of mitochondria drives mitochondrial network formation. Cell Res 25(10):1108–1120 4. Nunnari J, Suomalainen A (2012) Mitochondria: in sickness and in health. Cell 148(6): 1145–1159 5. Lewis SC, Uchiyama LF, Nunnari J (2016) ER-mitochondria contacts couple mtDNA

synthesis with mitochondrial division in human cells. Science 353(6296):aaf5549 6. McArthur K et al (2018) BAK/BAX macropores facilitate mitochondrial herniation and mtDNA efflux during apoptosis. Science 359(6378) 7. Qin J et al (2020) ER-mitochondria contacts promote mtDNA nucleoids active transportation via mitochondrial dynamic tubulation. Nat Commun 11(1):4471 8. Prole DL, Chinnery PF, Jones NS (2020) Visualizing, quantifying, and manipulating mitochondrial DNA in vivo. J Biol Chem 295(51): 17588–17601

Chapter 7 Visualization of mtDNA Using FISH Xie Xie and Xuefeng Zhu Abstract Proper mitochondrial DNA (mtDNA) levels are critical for many cellular biological functions and are associated with aging and many mitochondria disorders. Defects in core subunits of the mtDNA replication machinery lead to decreased mtDNA levels. Other indirect mitochondrial contexts including ATP concentration, lipid composition, and nucleotide composition also contribute to mtDNA maintenance. Furthermore, mtDNA molecules are distributed evenly throughout the mitochondrial network. This uniform distribution pattern is required for oxidative phosphorylation and ATP production and has been linked to many diseases when perturbed. Thus, it is important to visualize mtDNA in the cellular context. Here we provide detailed protocols for cellular visualization of mtDNA using fluorescence in situ hybridization (FISH). The fluorescent signals are targeted to the mtDNA sequence directly, ensuring both sensitivity and specificity. This mtDNA FISH method can be combined with immunostaining and used for visualizing mtDNA-protein interactions and dynamics. Key words mtDNA, FISH, Visualization, Microscopy, Mitochondria, Imaging

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Introduction The mammalian mitochondrial genome (mtDNA) is essential for cellular energy conversion, encoding 13 of the key subunits of the oxidative phosphorylation system (OXPHOS), as well as tRNA and rRNA molecules required for their translation [1, 2]. The mtDNA copy number varies depending on cell type and energy requirements, ranging from hundreds to many thousands per cell [3]. Maintaining proper mtDNA levels is critical for proper development and health [4], and defects in mtDNA maintenance cause a heterogeneous group of mitochondrial disorders. These mitochondrial disorders are usually characterized at the molecular level by mtDNA depletion, which leads to OXPHOS deficiency in affected tissues [2, 5]. The underlying causes of many of these disorders are mutations in the core subunits of the mtDNA replication machinery, including DNA polymerase γ (POLγ), the replicative DNA helicase

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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TWINKLE, and mtSSB [1]. Besides the replication factors, several indirect mechanisms are also proposed to affect mtDNA levels including ATP concentration, nucleotide composition, and mtDNA origin regulation [6]. Therefore, visualizing mtDNA at the cellular level could be used for looking for novel mtDNA regulators and provide new mechanistic insights on mtDNA maintenance. The mtDNA, which is packaged by TFAM, is located in the mitochondrial matrix and is well distributed throughout the mitochondrial network [7]. The distribution of mtDNA is important for the uniform distribution of mtDNA-encoded proteins in mitochondria and defects in mtDNA distribution are linked to many human diseases. Many biological processes, including mtDNA segregation, mitochondria fission and fusion, mitochondrial inner membrane structure, and lipid composition, can affect mtDNA distribution [8, 9]. Here we describe the protocols to visualize mtDNA using FISH, from cell seeding, probe labeling, to fixation and permeabilization, pre-hybridization, hybridization, combination with immunofluorescence staining, and final mounting for slides. Previous studies have used antibodies targeting either double-stranded DNA or mtDNA-associated proteins such as TFAM and TWINKLE to visualize mtDNA [10]. In comparison, mtDNA FISH uses sequence-specific probes to ensure specificity. In combination with FISH imaging analysis, it provides a quantitative measurement of mtDNA levels, especially in cells with low levels of mtDNA. This mtDNA FISH protocol can be used in super resolution microscopy, including STED and SIM, which is ideal for investigating mtDNA distribution and protein interactions.

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Materials

2.1 Cell Culture and Seeding

1. HeLa cells grown at 37 °C and 5% CO2. 2. DMEM GlutaMAX. 3. Gibco Fetal Bovine Serum (FBS). 4. Trypsin. 5. Sterile coverslips 1.5H 12 mm (Nordic Biolabs, Cat# 117520). 6. 24-well cell culture plates. 7. 100 mM 2′,3′-dideoxycytidine 5′-triphosphate (ddCTP) (Sigma, Cat# GE27-2061-01). 8. (Optional) For siRNA: Optimen (cold), Lipofectamine RNAiMAX, and siRNA (20 μM) of interest.

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mtDNA Probes Oligonucleotides

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1. h-mtF1 (FISH): 5′ ACAACCCCCGCCCATCCTACC 3′. 2. h-mtF2 (FISH): 5′ ACCTTCAAATTCCTCCCTGTACG 3′. 3. h-mtF3 (FISH): 5′ GGCAACCTTCTAGGTAACGACCA 3′. 4. h-mtF4 (FISH): 5′ GTAAGCCTCTACCTGCACGACAA 3′. 5. h-mtF5 (FISH): 5′ TCCATGCATTTGGTATTTTCGTC 3′. 6. h-mtR1 (FISH): 5′ CGAAGGGTTGTAGTAGCCCGTAG 3′. 7. h-mtR2 (FISH): 5′ ATGGCCCCTAAGATAGAGGAG 3′. 8. h-mtR3 (FISH): 5′ GAGGAGCGTTATGGAGTGGAAGT 3′. 9. h-mtR4 (FISH): 5′ GTGGATGCGACAATGGATTTTAC 3′. 10. h-mtR5 (FISH): 5′ TCCATGCATTTGGTATTTTCGTC 3′.

2.2.2 mtDNA Probe Preparation

1. Total cellular DNA from HeLa cells. 2. Taq DNA polymerase and buffer. 3. ChIP DNA Clean & Concentrator (Zymo, Cat# D5201). 4. Atto488 Nick Translation (NT) Labeling kit (Jena bioscience, Cat# PP-305 L-488). 5. Nanodrop 2000.

2.3 Chemicals and Equipment

1. PBS tablets, pH 7.4 (Medicago, Cat# 09-8912-100). 2. 16% (w/v) Formaldehyde, methanol-free (Thermo Fisher Scientific, Cat# 28908). 3. DAPI (Thermo Fisher Scientific, Cat# D1306). 4. Dextran sulfate (Merck, Cat# D8906). 5. Fixogum Rubber Cement (MP Biomedicals, Cat# FIXO0125). 6. Formamide (deionized) (see Note 1) (Thermo Fisher Scientific, Cat# AM9344). 7. RNase A, Thermo Fisher Scientific, Cat# EN0531. 8. RNase H, New England Biolabs, Cat# M0297. 9. DNase I set (Zymo, Cat# E1010). 10. 1 mg/mL Salmon sperm DNA (Merck, Cat# D7656). 11. Triton X-100 (Merck, Cat# T8787). 12. Tween-20 (Merck, Cat# P7949). 13. Parafilm. 14. Microscope slides. 15. Water bath. 16. Forceps. 17. BSA (Sigma, Cat# 10735086001). 18. Mounting medium.

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19. 0.2 micron filters. 20. Metal heating block. 2.4 Buffers and Solutions

1. 10× PBS, pH 7.4. Dissolve one PBS tablet in ddH2O, final volume 50 mL. Store at RT. 2. 10% Triton X-100. Measure out 10 mL Triton X-100, add ddH2O up to 100 mL. Store at 4 °C. 3. 0.5% Triton X-100/PBS: 0.5% Triton X-100 in 1× PBS. For 50 mL, combine 2.5 mL of 10% Triton X-100, 5 mL of 10× PBS, and 42.5 mL ddH2O. Store at RT. 4. PBS, pH 7.4. Dissolve one PBS tablet in ddH2O, final volume 500 mL. Store at RT. 5. RNase/PBS. Dilute RNase A (10 mg/mL) 1:100 and RNase H (5000 U/mL) 1:50 in 100 μL final volume PBS. 6. 20× SSC: 3 M NaCl, 0.3 M sodium citrate dihydrate. Dissolve 87.65 g of NaCl and 44.1 g of sodium citrate in 400 mL ddH2O. Adjust pH to 7.0 using 1 M HCl, add ddH2O to 500 mL. Filter using a 0.2 micron filter. Store at RT. 7. 2× SSC. Measure out 50 mL of 20× SSC, add ddH2O up to 500 mL. Store at RT. 8. 1× SSC/0.5× PBS. Mix together 50 mL of 2× SSC and 50 mL of PBS. 9. 10% Tween-20. Measure out 10 mL Tween-20, add ddH2O up to 100 mL. Store at 4 °C. 10. 4× SSCT: 4× SSC, 0.2% Tween-20. Combine 100 mL of 20× SSC and 10 mL of 10% Tween-20, bring volume up to 500 mL with ddH2O. Store at RT. 11. 2× SSCT/50% formamide (prepare just before use). Mix 4× SSCT with formamide at a 1:1 ratio. 12. 25% Dextran sulfate solution (see Note 2). Dissolve 5 g dextran sulfate in 16 mL ddH2O, store at -20 °C. 13. 2× Hybridization buffer: 20% dextran sulfate, 4× SSC. Combine 0.8 mL of 25% dextran sulfate and 0.2 mL of 20× SSC, store at RT. 14. Wash buffer (prepare fresh): 2× SSCT, 50% formamide, 0.1% Triton X-100. Combine 10 mL 4× SSCT, 10 mL formamide, and 0.2 mL 10% Triton X-100. 15. 2× SSCT: 2× SSC, 0.1% Tween-20. Combine 50 mL of 20× SSC and 5 mL of 10% Tween-20, bring volume up to 500 mL with ddH2O. Store at RT. 16. 0.2× SSC. Measure out 5 mL of 20× SSC, add ddH2O up to 500 mL. Store at RT.

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17. 2% BSA/PBST. Combine 2 mL 10% BSA, 1 mL 10× PBS, 0.1 mL 10% Tween-20, and 6.9 mL ddH2O. 18. 0.5% BSA/PBST: Combine 0.5 mL 10% BSA, 1 mL 10× PBS, 0.1 mL 10% Tween-20, and 8.4 mL ddH2O. 19. PBST: 0.1% Tween-20 in 1× PBS. For 50 mL, combine 0.5 mL of 10% Tween-20, 5 mL 10× PBS, and 44.5 mL ddH2O. Store at RT. 20. DAPI/PBS (1:50000). Dilute 1 μL DAPI in 50 mL final volume PBS. 21. 4% formaldehyde/PBS (make fresh). Combine 250 μL of 16% formaldehyde, 100 μL of 10× PBS, and 650 μL ddH2O.

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Methods

3.1 Cell Seeding (3 Days Before Staining)

1. Place sterile cover slips in a 24-well culture plate (one per well). 2. Trypsinize cells, measure cell concentration. 3. Dilute cells to 40,000/mL in DMEM GlutaMAX supplemented with FBS (10% final), seed 500 μL per well (see Note 3). 4. For ddCTP-treated samples, apply 0.5 μL ddCTP solution (concentration varies) to reach 0–10 μM final concentration (see Note 4). 5. If siRNA knockdown applies, mix together 1 mL optimen (cold), 8 μL lipofectamin RNAiMAX, 3 μL siRNA (20 μM), vortex and incubate at RT for 20 min. Add 83 μL mix per well and mix. 6. After 72 h incubation, cells are ready for fixation.

3.2 Preparing and Labeling mtDNA Probes

1. PCR amplify five mtDNA probes from the human mitochondrial genome using the oligonucleotide primers listed in Subheading 2.3, clean up reactions using the ChIP DNA Clean & Concentrator kit, and create a mix of the mtDNA probes (mtDNA mix) by pooling the PCR products (see Note 5). 2. Set up probe labeling reactions on ice, then mix briefly: 1 μg mtDNA mix 2 μL Atto488 NT labeling mix 2 μL NT labeling buffer 2 μL enzyme mix ddH2O up to 20 μL. 3. Incubate at 15 °C for 90 min using a PCR machine. 4. Add 30 μL ddH2O to the reaction, then add 250 μL DNA binding buffer from the ChIP DNA Clean & Concentrator kit.

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Fig. 1 Quality control of probe labeling by nick translation. (a) Representative absorbance profile of labeled probe using Nanodrop microarray analysis. A DNA peak is observed at 260 nm, and a second peak is observed at 501 nm, corresponding to the absorbance of Atto488. (b) Agarose gel showing size distribution of labeled probe after nick translation. The mtDNA template mix has a size around 3 kb, and after nick translation, the DNA smear size ranges from 100 to 300 bp

5. Load onto a DNA clean & concentrator column, centrifuge 30 s at top speed in a microcentrifuge. 6. Wash twice with 200 μL wash buffer, centrifuge 1 min at top speed. 7. Elute with 20 μL Elution buffer at top speed. 8. Measure the concentration and 488 incorporation using the Nanodrop microarray measurement (see Note 6) (Fig. 1). 3.3 Fixation and Permeabilization

1. Rinse cells twice with PBS. 2. Fix cell in 500 μL 4% formaldehyde/PBS at RT for 20 min. 3. Rinse twice with PBS. 4. Permeabilize with 500 μL 0.5% Triton X-100/PBS at RT for 10 min. 5. Wash 4× with 500 μL PBS for 1 min each.

3.4

RNase Treatment

1. Put 25 μL/sample RNase/PBS on parafilm, remove coverslips from the 24-well plates and place upside down onto the drop, incubate at 37 °C, 1 h, in a humidified chamber (see Note 7). 2. Transfer coverslips back into 24-well plates, wash 3× with 500 μL PBS for 1 min each. 3. (Optional) For samples needing DNase treatment, use the same setup as for the RNase treatment. Prepare DNase mix using 10 μL 10× buffer +10 μL DNase I + 80 μL ddH2O (see Note 8). Incubate at 37 °C, 1 h, in a humidified chamber. Wash 3× with 500 μL PBS for 1 min each. 4. Rinse cells with 500 μL 1× SSC/0.5× PBS. 5. Wash once with 500 μL 2× SSC.

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Prehybridization

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1. Add 500 μL 2× SSCT, incubate at RT, 5 min. 2. Add 500 μL of freshly prepared 2× SSCT/50% formamide, incubate at RT, 5 min. 3. Denature salmon sperm DNA at 99 °C for 5 min and then store on ice. 4. Place 8.7 μL of freshly prepared prehybridization buffer on a clean microscope slide. For example, for 8 coverslips, prepare a total of 160 μL prehybridization buffer: 80 μL 2× hybridization buffer +80 μL formamide +2.56 μL denatured ssDNA (16 ng/ μL final). 5. Transfer coverslips on top of the prehybridization microscope slide (see Note 9). 6. Seal the coverslip onto the slide by adding a layer of rubber cement, completely covering the coverslip. Allow rubber cement to dry for 20 min. 7. Set up a heating block at 92 °C, reverse block (bottom side up). Denature the sample at 92 °C by placing the slide onto the block for 2.5 min. 8. Place slide in a humidified chamber at 37 °C, 1.5 h.

3.6

Hybridization

1. Add mtDNA probe (from Subheading 3.2) to new prehybridization buffer at a final concentration of 2 ng/μL (see Note 10). 8.7 μL of the mixture is needed per coverslip, scale up as needed. 2. Per sample, put 8.7 μL hybridization buffer + probe onto a clean microscope slide. 3. Transfer coverslips, from step 7 in Subheading 3.5, on top of the hybridization microscope slide, cells facing down (no pressing). 4. Seal the coverslip onto the slide by adding a layer of rubber cement, completely covering the coverslip. Allow rubber cement to dry for 20 min. 5. Denature at 92 °C for 2.5 min, as in step 7 in Subheading 3.5. 6. Transfer to a humidified chamber and incubate at 37 °C overnight.

3.7

Washes

1. Pre-warm the wash buffer at 37 °C in a water bath. 2. Carefully remove the coverslip from the glass slides with forceps and put in 24 well plate, cells facing up. All wash steps use 500 μL volume per well. 3. Wash with pre-warmed wash buffer at 37 °C for 15 min, three times. 4. Wash with wash buffer at RT for 15 min, twice.

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5. Wash with 2× SSCT at RT for 15 min (with gentle shaking). 6. Wash with 0.2× SSC at RT for 10 min (with gentle shaking). 7. Wash with PBS at RT for 5 min. 3.8 Immunostaining (Optional) and Nuclear Counterstaining (see Note 11)

For the following steps, place 50 μL drops of the solutions on a piece of Parafilm, and place each coverslip cell side down on the separate drops on the Parafilm. 1. Block in 2% BSA/PBST for 1 h, RT. 2. Incubate with primary antibody, diluted 1:200 in 0.5% BSA/PBST, 1 h, RT. 3. Wash with PBST at RT for 2 min, three times. 4. Incubate with secondary antibody, diluted 1:250 in 0.5% BSA/PBST, 1 h, RT. 5. Wash with PBST at RT for 2 min, three times. 6. Add DAPI/PBS stain for 5 min. 7. Wash with PBS for 2 min, two times.

3.9

Mount

1. Apply 5 μL of mounting medium to the surface of a slide. 2. Remove coverslip containing the sample from the buffer. 3. Blot excess buffer from the non-sample surface of the coverslip. 4. Slowly tip the coverslip onto the mounting medium and avoid creating bubbles as you lower it into place. 5. Store the coverslip at room temperature overnight in the dark to cure the samples. 6. Slides are ready for imaging using a fluorescent microscope.

4

Notes 1. Formamide used for pre-hybridizaiton solution and hybridization solution are stored as frozen aliquots (freshly deionized). Formamide used in all washes is stored at 4 °C. 2. Dissolve dextran sulfate in water overnight at 4 °C, rocking, or warm up to 65 °C. Add ddH2O to 20 mL, make 0.8 mL aliquots and freeze at -20 °C. 3. Dilute cells to final concentration in 50 mL Falcon tube. Shake the tube mildly. After adding cells to the wells, swirl the plate gently to make sure cells are distributed evenly on the plate. 4. Treatment with ddCTP results in a decrease of mtDNA. Make four serial concentrations (0, 2.5, 5, and 10 μM) to obtain different levels of mtDNA (Fig. 2).

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Fig. 2 ddC treatment results in decreasing level of mtDNA. Representative images of mtDNA FISH in Hela cells treated with different concentrations of ddC. Scale bar, 20 μm

5. We designed five primer pairs to amplify the human mtDNA genome (except the D-Loop region). Each primer pair amplifies about 3 kb. The standard PCR protocol using Taq polymerase and total DNA from Hela cells works well for amplification. Equal amounts of the five PCR products are pooled to obtain a mix of mtDNA probes which covers most of the human mitochondrial genome. 6. In Nanodrop ND2000, choose the Micro array tab. For sample type, choose DNA. For concentration unit, choose ng/μL. For dye1, choose Alexa Fluor 488, and for the concentration unit choose pmol/μL. A successful labeling can produce more than 0.1 pmol of 488 incorporation per ng of DNA, e.g., 80 ng/μL DNA with >8 pmol/μL 488 incorporation. 7. Make a humidifying chamber using wet paper in a sealed container to prevent evaporation. 8. DNase treatment can completely remove DNA content; it should be performed regularly to avoid false contamination fluorescent signals (Fig. 3). 9. Do not press the coverslip by force. Up to four coverslips can be placed on the same coverslip. 10. From here on, keep the probe and coverslip in the dark. 11. This FISH protocol can be combined with immunostaining, such as for the mitochondrial marker COX IV. However, some antibodies, such as anti-TFAM, cannot be used due to the high denaturing temperature. Individual tests for each antibody are required.

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Fig. 3 Representative confocal microscopy images of mtDNA FISH in HeLa cell subjected to different nuclease treatments. Please note that mtDNA FISH probes also stain mtRNA and produce modest signal. Thus, an RNase treatment step is required to ensure specificity of mtDNA signal. Scale bar, 5 μm References 1. Gustafsson CM, Falkenberg M, Larsson NG (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160 2. Viscomi C, Zeviani M (2017) MtDNAmaintenance defects: syndromes and genes. J Inherit Metab Dis 40:587–599 3. Reznik E et al (2016) Mitochondrial DNA copy number variation across human cancers. elife 5 4. Moraes CT (2001) What regulates mitochondrial DNA copy number in animal cells? Trends Genet 17:199–205 5. Clay Montier LL, Deng JJ, Bai Y (2009) Number matters: control of mammalian mitochondrial DNA copy number. J Genet Genomics 36:125–131 6. Filograna R, Mennuni M, Alsina D, Larsson NG (2021) Mitochondrial DNA copy number

in human disease: the more the better? FEBS Lett 595:976–1002 7. Campbell CT, Kolesar JE, Kaufman BA (2012) Mitochondrial transcription factor a regulates mitochondrial transcription initiation, DNA packaging, and genome copy number. Biochim Biophys Acta 1819:921–929 8. Stefano GB, Kream RM (2016) Mitochondrial DNA heteroplasmy in human health and disease. Biomed Rep 4:259–262 9. Stewart JB, Chinnery PF (2015) The dynamics of mitochondrial DNA heteroplasmy: implications for human health and disease. Nat Rev Genet 16:530–542 10. Kukat C et al (2011) Super-resolution microscopy reveals that mammalian mitochondrial nucleoids have a uniform size and frequently contain a single copy of mtDNA. Proc Natl Acad Sci U S A 108:13534–13539

Chapter 8 In Situ Analysis of Mitochondrial DNA Synthesis Using Metabolic Labeling Coupled to Fluorescence Microscopy John A. Smolka and Samantha C. Lewis Abstract Metabolic labeling with the nucleoside analog 5-ethynyl-2′-deoxyuridine (EdU) enables the selective labeling of DNA synthesis in live cells. Newly synthesized EdU-containing DNA can be covalently modified after extraction or in fixed cells using copper-catalyzed azide-alkyne cycloaddition “click chemistry” reactions, enabling bioconjugation to various substrates including fluorophores for imaging studies. While often used to study nuclear DNA replication, EdU labeling can also be leveraged to detect the synthesis of organellar DNA in the cytoplasm of Eukaryotic cells. In this chapter, we outline methods for the application of EdU labeling to the study of mitochondrial genome synthesis in fixed cultured human cells, using fluorescent labeling and superresolution light microscopy. Key words Metabolic labeling, EdU, Mitochondrial DNA, Click chemistry, Superresolution microscopy

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Introduction Mitochondria house and propagate a genome separate from the nuclear genome, referred to as mitochondrial DNA (mtDNA). Though relatively small (~16,000 base pairs), human mitochondrial genomes encode genes for a mitochondrial protein expression system (ribosomal RNA, transfer RNAs, and key electron transport chain proteins) essential for cellular respiration. Individual mammalian cells can contain hundreds to thousands of copies of mtDNA [1]. The propagation and abundance of mtDNA requires at least 4 proteins encoded in the nucleus: POLG1, POLG2, SSBP1, and TWNK, which are thought to constitute a minimal mitochondrial replisome in humans [2]. Defects in mtDNA replication, along with depletion and/or mutation of mtDNA, are etiological factors in human mitochondrial diseases [3]. Unfortunately, our current understanding of mitochondrial genome

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maintenance is impeded by a lack of high-fidelity, high-resolution, and quantitative readouts of mtDNA replication in cells. Prior methods, such as quantitative polymerase chain reaction (qPCR) and 2-dimensional neutral agarose gel analyses, do not directly assay DNA synthesis. Additionally, these methods yield aggregate measurements of mtDNA abundance and structural features derived from populations of cells and thus cannot provide information on the order of a single cell, single mitochondrion, nor single mitochondrial genome. Another method, DNA combing/ fiber analyses, using pulses of nucleoside analogs in live cells followed by DNA extraction and fluorescent labeling, can provide information on the kinetics of mtDNA replication at single genome resolution, yet is also inherently destructive to cells [4]. Altogether, these approaches provide a diverse and informative toolkit for measuring mtDNA abundance, structure, and replication kinetics, but also require DNA extraction from cells and thus lack cytological and cellular context. Metabolic labeling via nucleoside analogs followed by fluorophore conjugation of DNA in fixed cells enables the visualization of mtDNA replication in cellular context, as well as a quantitative readout for mtDNA synthesis at single cell, single organelle, and potentially single genome, resolution. Thus, in addition to measuring the levels of mtDNA synthesis in individual cells, the cytological distribution of mtDNA replication initiation and potential spatial links with organellar structure and other protein factors can be assessed [5]. This can be accomplished using antibody-based detection of a nucleoside analog, for example Bromodeoxyuridine (BrdU), or via covalent modification of EdU with a fluorophore via click chemistry. Immunolabeling of incorporated nucleoside analogs requires access to the modified base by an antibody, which, for assaying DNA replication in situ, typically entails acid treatment to partially hydrolyze DNA and facilitate denaturation of double-stranded DNA prior to immunolabeling. The use of EdU and click chemistry obviates the need for such steps, presumably because the fluorescent azides used are small enough to access the ethynyl group of EdU in the context of a native DNA duplex [6]. Here, we provide a standardized EdU labeling protocol, with controls and assays for reagent validation, useful for studying mitochondrial DNA synthesis in situ. Using super-resolution microscopy, we demonstrate that fluorescently conjugated EdU can be imaged in tandem with immunolabeled mitochondrial DNA and mitochondrial membrane(s) to obtain high-resolution information on the spatial distribution of mitochondrial genome synthesis (Fig. 1). These data permit the dissection of heterogeneity in the regulation of mtDNA synthesis amongst the mitochondrial genomes within single cells and the comparative assessment of global mtDNA replication levels between individual cells.

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Fig. 1 (a) A max intensity projection (MIP) of a fixed U2OS cell metabolically labeled with a 1 h pulse of 10 μM EdU followed by a 1 h chase, and immunolabeled with anti-dsDNA and anti-TOMM20 antibodies. Scale bar: 5 μm. (b) MIPs of cells from additional U2OS samples prepared identically to and in parallel with the sample from (a). Cells were treated postfixation/prior to click reactions and immunolabeling with either benzonase or RNase A for 1 h at room temperature to assess the DNA dependence of both EdU and anti-dsDNA antibody signals (see Note 4 for treatment protocol)

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Materials Reagents

1. DMEM + glutaMAX, with 4.5 g/L glucose (10566-016, Gibco). 2. Click-iT Plus EdU Alexa Flour 647 Imaging Kit (C10640, Invitrogen). 3. Glass bottom poly-D-lysine-coated dishes (P35GC-1.5-14-C, MatTek). 4. 4% (v/v) paraformaldehyde in PBS. 5. PBS with 0.1% (v/v) Triton X-100. 6. PBS with 0.1% (v/v) Tween and 1% (w/v) BSA. 7. Benzonase (E8263-5KU, Sigma-Aldrich). 8. RNase A (EN0531, ThermoFisher). 9. Rabbit anti-TOMM20 (Proteintech, 11802-1-AP). 10. Mouse anti-dsDNA (Abcam, ab27156) 11. AlexaFluor 405 goat anti-mouse (A31553, Invitrogen). 12. AlexaFluor 488 donkey anti-rabbit (A32790, Invitrogen).

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Equipment

1. Zeiss 980 LSM Airyscan II microscope on inverted stand equipped with Plan-Apochromat 63×/1.4NA oil objective and ZEN software 3.0.

Methods

3.1 EdU Pulse Labeling

1. Plate cells at ~20% density (~20,000–30,000 cells) in a 35 mm glass bottom poly-D-lysine-coated dish; for this experiment, 2 mL media volumes were used for each sample. 2. Grow cells for 24–48 h in a humidified cell culture incubator at 37 °C with an atmosphere of 5% carbon dioxide, until they reach 40–60% confluency. 3. Aliquot 1 uL of 10 mM EdU stock to a microfuge tube. Aliquot 1 mL of media from the sample to be labeled to a second microfuge tube. Remove 0.5 mL of the remaining media in the dish and add this directly to the microfuge tube containing 1 uL of EdU and mix briefly by pipetting up and down. Immediately add the media/EdU mix back to the glass bottom dish sample (final EdU concentration: 10 uM), and briefly mix by gently shaking or rocking the sample. 4. Incubate both the samples and saved media for the desired pulse time in a cell incubator. 5. Thoroughly remove EdU pulse media by vacuum suction and replace it with the preserved, conditioned EdU-free media. Incubate for an additional 15 min (this is the chase time), or longer if desired, in the incubator.

3.2 Fixation and Click Reaction

1. Fix cells by removing the culture medium by vacuum suction and adding 2 mL of 4% paraformaldehyde diluted in PBS (pre-warmed to 37 °C) and then incubate the sample for 20 min at room temperature. *All steps from here on are performed at room temperature. 2. Quench fixation solution by adding 200 uL of 1 M glycine in PBS directly to the fixation reaction/fixed sample. Mix by gentle rocking/shaking and incubate for 5 min. 3. Remove quenched fixation solution, and gently wash sample once with PBS. 4. Add 2 mL of permeabilization solution, gently mix sample, and incubate for 10 min. 5. Remove permeabilization solution, add 2 mL of staining/ blocking buffer, and incubate for 10 min. 6. Remove staining/blocking buffer, and add 0.5 mL of complete click reaction cocktail with azide (consisting of 440 uL of

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reaction buffer, 10 uL of copper protectant solution, 1.2 uL of fluorescent azide, and 50 uL of click reaction additive), applying dropwise to the side of the dish (not directly on the glass bottom of the dish/cells to be imaged) (see Notes 1 and 2). Gently tap the dish several times until the glass is completely covered by the reaction cocktail. Gently shake/rock the sample while incubating for 30 min, protected from light. *Sample(s) should be protected from light when possible for all subsequent steps. 7. Add 1.5 mL of staining/blocking buffer to the sample, mix, and then remove the 2 mL of diluted reaction cocktail. Wash the sample once with staining/blocking buffer. Sample can then be immunolabeled/prepared for imaging using a protocol of choice, or the following immunolabeling protocol. 3.3

Immunolabeling

1. Add 2 mL of primary antibody dilution (1:1000 dilution for all antibodies used here) made in staining/blocking buffer. Incubate at room temperature for a minimum of 1 h, or overnight at 4 °C. 2. Wash once with 2 mL of staining/blocking buffer, and then apply secondary antibodies (typically at a 1:2000 dilution) in the same manner used for primary labeling. Incubate for 1 h at room temperature. DNA stains, like DAPI or Hoechst diluted in blocking/labeling solution, can be applied after this step, if desired; we recommend using the Hoechst stain provided with the Click-iT kit at a 1:2000 dilution with a 5 min incubation at room temperature. 3. Wash once with a 10 min incubation in 2 mL staining/blocking buffer. 4. If imaging within 48 h (recommended): remove wash and add 2 mL of PBS. Samples can be stored at 4 °C protected from light; bring dishes to room temperature before imaging. Imaging can be performed with no further mounting medium. 5. If not imaging within 48 h: remove PBS thoroughly by vacuum suction and then add two drops of Prolong Diamond directly to the center of the dish to permanently preserve the sample in hardening mounting medium. Let sample cure for 48 h, protected from light prior to imaging.

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Imaging

1. To image fixed adherent mammalian cells such as the osteosarcoma cells used here, we recommend the Zeiss 980 LSM Airyscan II microscope on an inverted stand, selecting the PlanApochromat 63×/1.4NA oil objective. 2. To set up the acquisition software, open Zeiss ZEN version 3.0 or higher and select the following parameters: zoom factor 1.5

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×; maximum pixel dwell time 0.83 μs/pixel; pixel size 35 nm/ pixel (2 × Nyquist optimal); environmental chamber 22 °C without supplemental humidity. 3. Align the light path/Airyscan detectors by selecting the “Adjust in live and continuous scans” setting under the Airyscan Detector Alignment menu, and then activate Continuous mode with SR-4Y multiline scanning, until the formerly detector icon turns green, indicating successful alignment. 4. For Z-stack acquisition, select a 4 μm total stack depth to capture an entire adherent mammalian cell in Z. The optimal step size for stack acquisition will vary depending on the sample; 0.15 μm was used for the images shown here. 5. 3D Airyscan process the acquired Z stacks in ZEN 3.0 by selecting “Airyscan Processing” from the Image Processing menu tab while the “3D” box is checked, then transform processed images into maximum intensity projections by selecting “Extended Depth of Focus” and “Max Intensity” in the ZEN 3.0 dialogue menu. (see Notes 3, 4, 5, and 6 for additional information about interpretation of fluorescence signals and caveats about detection of mitochondrial EdU).

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Notes 1. It is critical to strictly follow the protocol for making the click reaction cocktail provided by the Click-iT Plus EdU Alexa Flour 647 Imaging Kit. Specifically, click additive (ascorbate) must be added last, and the reaction mixture should always be made fresh and promptly applied to samples. Deviation from the prescribed order of reagent addition or incubation of the reaction mixture in excess of the recommended time risks compromised reaction efficiency. Ensuring maximum assay sensitivity is particularly important for robustly detecting DNA synthesis associated with 16 kb human mitochondrial genomes. 2. Copper solutions can damage DNA. Applying the click reaction cocktail directly to cells on glass risks exacerbating DNA degradation by the reaction mixture. We recommend applying the reaction mixture to the side of the dish or slide and then tilting to wash the solution over the fixed cells. The DNA damaging capacity of copper solutions should be considered for the smaller and potentially more fragile mitochondrial genome, and copper solutions used should always be supplemented with a chelating agent, as in the kit used in the protocol, to mitigate this potential source of damage.

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3. We find that mitochondrial EdU signal does not always overlap/colocalize precisely with anti-dsDNA signal. However, controls demonstrating sensitivity of both EdU and antidsDNA signals to benzonase and resistance to RNase A treatments support that the EdU signal stems from DNA polymers that are not labeled by anti-dsDNA. 4. We find that a 1 h treatment with benzonase at room temperature is sufficient to degrade the majority of mtDNA-associated signals in situ. However, we note that it did not remove the nuclear anti-dsDNA or nuclear EdU signal. We recommend benzonase and RNase A treatments be carried out for at least 1 h in 1 mL volumes with 1:500 dilutions of enzyme in a 1: 2 dilution of blocking/labeling solution (0.5 × TBS, 0.05% (v/v) Tween, 0.5% (w/v) BSA), followed by a wash with blocking/labeling solution. 5. Nuclear EdU signal is typically more intense than mtDNAassociated EdU signal. When imaging S-phase cells, the intensity of EdU labeled nuclei may potentially impede imaging of the cytoplasm. Using a Zeiss Airyscan 980, we have been able to image the cytoplasm of cells in S-phase with robust signalto-noise ratio. However, it is important to determine to what extent a user’s microscope system generates images affected by this issue. If noise is an issue, we recommend imaging only non-S-phase cells, and/or using aphidicolin pretreatments to inhibit nuclear DNA replication (see [5] for aphidicolin treatment protocol). 6. We have varied EdU pulse times, reducing them to 15 min, and been able to detect decreases and increases in mitochondrial EdU signal that are proportional to pulse times. However, to start, we recommend using a 1 h pulse time, which should provide robust labeling to use as a reference point for optimization.

Acknowledgments This work was supported by National Institutes of Health grant GM129456. References 1. Kelly RD, Mahmud A, McKenzie M, Trounce IA, St John JC (2012) Mitochondrial DNA copy number is regulated in a tissue specific manner by DNA methylation of the nuclear-encoded DNA polymerase gamma A. Nucleic Acids Res 40(20):10124–10138. https://doi.org/10. 1093/nar/gks770

2. Korhonen JA, Pham XH, Pellegrini M, Falkenberg M (2004) Reconstitution of a minimal mtDNA replisome in vitro. EMBO J 23(12): 2423–2429. https://doi.org/10.1038/sj. emboj.7600257 3. Copeland WC (2014) Defects of mitochondrial DNA replication. J Child Neurol 29(9):

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1216–1224. https://doi.org/10.1177/ 0883073814537380 4. Phillips AF, Millet AR, Tigano M, Dubois SM, Crimmins H, Babin L, Charpentier M, Piganeau M, Brunet E, Sfeir A (2017) Singlemolecule analysis of mtDNA replication uncovers the basis of the common deletion. Mol Cell 65(3):527–538 e526. https://doi. org/10.1016/j.molcel.2016.12.014

5. Lewis SC, Uchiyama LF, Nunnari J (2016) ER-mitochondria contacts couple mtDNA synthesis with mitochondrial division in human cells. Science 353(6296):aaf5549. https://doi. org/10.1126/science.aaf5549 6. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci U S A 105(7): 2415–2420. https://doi.org/10.1073/pnas. 0712168105

Chapter 9 Measurement of Nucleoid Size Using STED Microscopy Elisa Motori Abstract Mitochondria are equipped with their own DNA (mtDNA), which is packed into structures termed nucleoids. While nucleoids can be visualized in situ by fluorescence microscopy, the advent of superresolution microscopy, and in particular of stimulated emission depletion (STED), has recently enabled the visualization of nucleoids at sub-diffraction resolution. Super-resolution microscopy has proved an invaluable tool for addressing fundamental questions in mitochondrial biology. In this chapter I describe how to achieve efficient labeling of mtDNA and how to quantify nucleoid diameter using an automated approach in fixed cultured cells by STED microscopy. Key words Mitochondrial DNA, Nucleoid, Stimulated emission depletion microscopy, STED, Fluorescence microscopy, Immunocytochemistry, ImageJ

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Introduction Mitochondria are double membrane-bound organelles located in the cytoplasm of cells, which use oxidative phosphorylation (OXPHOS) to generate most of the energy supply for the cell in the form of ATP. Often nicknamed as the ‘powerhouse of the cell’, these organelles can perform a variety of metabolic processes beyond ATP production [1] and also contribute to important cellular processes such as regulation of calcium homeostasis [2] and apoptosis [3]. Notably, mammalian mitochondria contain their own DNA (mtDNA), a circular genome that encodes 13 proteins with essential roles in maintaining a functional OXPHOS system and whose altered expression is invariably associated with human diseases [4, 5]. Thus, understanding which mechanisms control mtDNA organization, distribution, and function within tissues is a fundamental question in mitochondrial biology with key implications for therapeutic approaches. Each cell can possess thousands of copies of mtDNA in the form of nucleoids, each usually containing a single mtDNA molecule compacted via cross-strand binding to its main protein

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constituent, mitochondrial transcription factor A (TFAM) [6]. By immunofluorescence, nucleoids appear as punctate foci spread across the mitochondrial network. Initial studies using confocal microscopy reported a mean size of nucleoids very close to the diffraction limit, suggesting that these structures could contain several copies of mtDNA [7, 8]. However, this interpretation was bound to be reassessed given the limited spatial resolution inherent in confocal microscopy. The development of STED (and related super-resolution microscopy techniques) has made it possible to review and expand on important aspects of nucleoid biology by breaking the diffraction limit to enable imaging of samples at ~20 nm resolution [9, 10]. In particular, recent studies not only revealed a significantly smaller mean nucleoid size than the one measured with conventional confocal microscopy [8, 11], but also provided a wealth of new information on nucleoid distribution under physiological conditions [12] or in cells with mitochondrial dysfunction [13, 14]. Although STED is typically built on a confocal system, it utilizes a double laser design: a standard excitation laser beam coupled with a second beam that has a doughnut shape (no intensity in the focal center but strong intensity in the periphery). As a result, excited fluorophores exposed to the doughnut laser return immediately to the ground state, while those located in the focal center can still emit fluorescence, thus creating a much smaller observed volume [15]. In this chapter, I describe how to quantify nucleoid size in STED images using a freely available software. This includes a protocol for sample preparation that meets the standards required by STED imaging.

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2.1 Immunocytochemistry

1. #1.5 or #1.5H (high performance) glass coverslips of the desired diameter (12 mm diameter coverslips provide enough surface for most cell types). Before use, wash coverslips in distilled water to remove any dirt or dust residue, then autoclave them (see Note 1). 2. Multiwell plates or dishes suitable for the diameter of the coverslips. 3. 1× Phosphate buffered saline (PBS), pH 7.4. 4. Permeabilization solution: Triton X-100 0.1% (v/v) in 1× PBS, to be stored at 4 °C. 5. Blocking buffer: bovine serum albumin (BSA) 3% (w/v) in 1× PBS. To prevent microbial growth, filter the solution with a

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0.22 μm Stericup® Vacuum filter (or similar) and prepare 50 mL aliquots. Store the solution at 4 °C. 6. Paraformaldehyde (PFA) 4% (v/v) in PBS, pre-equilibrated at room temperature (RT) (see Note 2). 7. Staining cassette. To prepare one, wrap a polystyrene square dish and its lid with aluminium foil, then place a layer of hydrophobic parafilm inside the dish (see Note 3). 8. Antibodies to visualize mitochondrial DNA. The protocol here described has been optimized using a mouse monoclonal IgM anti-DNA (clone AC-30-10; Progen, 1:200 dilution) and a goat anti-mouse IgM Alexa Fluor 594 (Thermo Fisher, 1: 2000 dilution) (see Note 4). 9. Mounting Medium. Aqua Polymount or Prolong Diamond are recommended. 2.2 Analysis of Nucleoid Diameter

3

1. ImageJ software (freely available at [16]), or Fiji. 2. Additional plug-ins (not included in the basic version of ImageJ): Adrian’s FWHM plugin (freely available at [17]).

Methods

3.1 Immunocytochemistry

Efficient labeling of the sample and structure preservation are important requirements in conventional confocal microscopy, but they become essential when considering the increased resolution achieved with STED microscopy. Here, a protocol for labeling mtDNA in fixed culture cells is provided that has been optimized for downstream STED imaging (see Note 5). This labeling protocol can be easily performed using standard equipment available in all wet labs. Unless otherwise stated, all the incubation steps should be carried out at room temperature and with the cassette closed to prevent bleaching of the fluorescent signal. The volumes indicated here are for 12 mm diameter coverslips and need to be scaled up or down according to the desired format. Imaging of the samples can be performed using a custombuilt STED microscope, or a commercially available one (see Note 6). Figure 1a, b shows the typical confocal and STED images of nucleoids in mouse embryonic fibroblasts labeled using this protocol. 1. Grow cells onto coverslips inside a multiwell plate for at least 24 h before fixation (see Note 7). After 24 h, or at the desired time if the experiment requires a treatment, take the cells out of the incubator and remove the medium. 2. Add 70 μL of fixative solution for 10 min at RT. 3. Remove the PFA (see Note 8).

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Fig. 1 Labeling of mitochondrial nucleoids for downstream STED imaging. (a) Representative confocal pictures showing nucleoids (grey puncta) of mouse embryonic fibroblast labeled as described in 3.1. The outer mitochondrial membrane marker Tom20 (red) is used to visualize mitochondria. A zoom of the cyan boxed region is provided below. Scale bars: 15 and 4 μm. (b) Confocal (left) and STED (right) images of the yellow boxed region in (a). The nucleoids that appear partially resolved in the confocal picture are completely discernible using STED. Scale bar: 500 nm. (c) Intensity profile analysis of the nucleoid in (b); the arrows indicate the FWHM obtained in confocal (dashed line) or STED (solid line) modality

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4. Add 70 μL of Permeabilization Buffer and incubate for 5 min at RT. 5. Remove the Permeabilization Buffer and add 70 μL 1× PBS to each well. With the help of forceps, transfer the coverslips from the multiwell plate to the staining cassette. Coverslips must be placed on the parafilm layer with the cells facing up. 6. Add 70 μL of Blocking Buffer. 7. In the meantime, prepare a master mix solution containing the primary antibody in 70 μL Blocking Buffer. 8. Remove the Blocking Buffer and add the solution containing the primary antibody to the slides. Close the cassette and incubate overnight at 4 °C. 9. Rinse three times with 100 μL Blocking Buffer for 5 min each time (see Note 9). Prepare a master mix solution containing the secondary antibody in 70 μL Blocking Buffer and add it to the slides. Close the cassette and incubate for 2 h at RT in the dark. 10. Quickly rinse the coverslips with 100 μL 1× PBS, then wash three times in 100 μL 1× PBS for 10 min each (see Note 10). 11. At the end of the washing steps, remove the PBS by placing the slide vertically onto a tissue. Apply one drop of mounting medium onto the glass cover and immediately mount the slide. Allow medium to cure overnight at RT. Store slides at 4 °C until imaging (see Note 11). 3.2 Nucleoid Analysis

A statistically robust and validated method to measure nucleoids diameter is based on the Full-Width at Half-Maximum (FWHM) function [8, 11]. Using this method, the intensity of the fluorescence signal of a given nucleoid is projected and fitted into a curved function, and the nucleoid diameter is calculated as the distance between the points where the intensity is half of the maximum (Fig. 1c). The FWHM can be determined either manually, or by using specific software-programmed scripts. While manual determination is time-consuming and prone to biased interpretation, automated scripts require software specific knowledge and the software itself may not be universally available. Here, an automated method that can be run on the opensource software ImageJ (or Fiji) is detailed. The method takes advantage of the Adrian’s FWHM plug-in, which was written by Adrian Martin to analyze photon detector pinhole images [17]. The plug-in is equipped with the ‘single pinhole’ function, which allows analysis of one event (nucleoid) at time; it generates more data than bulk analysis and may be recommendable when running the analysis for the first time on a dataset in order to check the suitability of the default settings. Further details on the operation and the algorithms used by the plug-in can be found in the code documentation at [17].

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3.2.1 Single Nucleoid Analysis

1. Open ImageJ and upload the image to be analyzed (see Note 12). Adjust the brightness and contrast. 2. Using the ROI manager function, outline a single nucleoid in the image. Avoid choosing nucleoids that are out of focus as this can affect the measurement. 3. Select ‘Adrian’s FWHM’ from the menu. Turn on the ‘Single pinhole’ button in the Multi Pinhole Search Option window (Fig. 2a, red dashed square). By selecting the ‘Advanced’ button (Fig. 2a, solid red square), a second window showing the advanced functions will pop up (Fig. 2a, red square; Fig. 2b): make sure the options ‘zoom of pinhole’ and ‘show fits’ are turned on if you need this information (see Note 13). 4. Press ‘OK’ to run the analysis. An example of the results table is shown in Fig. 3a with FWHM values for both the x- and y-axes (see Note 14). If the related options were selected before running the analysis, additional data such as the zoom of the analyzed pinhole and the histogram of the fit will also be shown (Fig. 3b). 5. Convert FWHM values into nm values using the formula: ½FWHM∙½pixel size

Fig. 2 Overview of Adrian’s FWHM plug-in. (a) Screenshot of a STED image of nucleoids being analyzed with the Adrian’s FWHM plug-in. When using the ‘single pinhole’ function, the nucleoid of interest is outlined (yellow circle) with the selection function in the toolbar, and the single pinhole function is activated in the Multi-Pinhole search option window. Advanced settings are shown in (b)

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Fig. 3 Nucleoid analysis with the single pinhole function. (a) Results window showing the FWHM values for the single nucleoid outlined in yellow. The Gaussian fit and the box with the zoom corresponding to the analyzed nucleoid are shown in (b)

Express the diameter as the average of FWHM measurements of the x- and y-axis (see Note 15). 3.2.2

Bulk Analysis

1. Open ImageJ and upload the image to be analyzed (see Note 12). Adjust the brightness and contrast. 2. Using the ROI manager function, outline a region of interest in the image (see Note 16). Avoid choosing nucleoids that are out of focus as this can affect the measurement. 3. Select ‘Adrian’s FWHM’ from the menu. 4. Press OK to run the analysis. An example of the results table is shown in Fig. 4 with FWHM values for both the x- and y-axes (see Note 14). 5. Convert the FWHM values into nm values using the formula: ½FWHM∙½pixel size Express the diameter as the average of FWHM measurements of the x- and y-axis (see Note 14).

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Fig. 4 Bulk analysis of nucleoids. (a) The nucleoids of interest are outlined (yellow ellipse) with the selection function in the toolbar, and the corresponding FWHM values are shown in the Results window on the right

4

Notes 1. The microscope objective lens used in confocal imaging is usually corrected for glass coverslips with a thickness of 0.17 mm, which is very close to that of #1.5 (or #1.5H) coverslips. Using the incorrect coverslip thickness can significantly affect the resolution by introducing spherical aberrations, particularly in the case of super-resolution techniques. 2. Fixation is a critical step, since it determines how much the mitochondrial (and nucleoid structure) will be preserved in the sample. For optimal results, it is recommended to prepare a fresh solution each time and to use high quality PFA (such as EM grade) that is devoid of methanol, a chemical that can distort mitochondrial morphology. I routinely dilute a 16% PFA EM grade aqueous solution (Electron Microscopy Sciences) with 1.3× PBS and store it at 4 °C until use. Since the PFA stock is an aqueous solution, 1.3× PBS is here used to compensate for osmolality. 3. To prevent drying of the slides, put a folded piece of paper tissue previously soaked in water in one corner of the cassette, far from the coverslips.

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4. The labeling steps described in this protocol have been optimized using the antibodies listed, and so may require further optimization when using different antibodies or specimens. Also, if the experiment requires multicolor STED, please consult the following source for suitable fluorophores combinations [18]. 5. Providing a comprehensive guide to sample preparation is beyond the scope of this chapter, and further details on sample preparation and suitable fluorophores for STED can be found at [18, 19]. 6. Several aspects should be taken into account to ensure optimal imaging conditions for STED, such as the available laser technology, objectives, and scanning parameters [15]. These aspects greatly differ and should be optimized depending on the available STED microscope; a systematic discussion of these aspects can be found here [20]. Also, the instrument performance should be tested before running an experiment by employing custom-made bead samples (a protocol to prepare bead samples can be found here [19]) or commercially available nanorulers (e.g., from GATTAquant). 7. Although cells attach and start to spread on the glass within hours, most cells need 1–2 days to reacquire normal morphology. To preserve the morphological and structural features of the cells and ensure a proper labeling density, it is important to keep culture subconfluent with cells spread flat. 8. If the staining protocol cannot be started the same day, the procedure can be stopped here and coverslips can be stored in 1× PBS at 4 °C. However, it is recommended to stain the coverslips within a couple of days after fixation, since prolonged storage of fixed cells will significantly affect the quality of the staining. 9. A larger volume of solution is applied during the washing steps to ensure efficient removal of unbound antibody. 10. The quick rinsing step is done to remove the excess unbound secondary antibody, which can add to the background noise. Longer and/or more washing steps can be performed to increase the quality of the labeling. 11. Follow the manufacturer’s instruction on the curing time of hardening mounting media. Although slides can be stored at 4 °C, it is recommended to image samples within a couple of days for the best results. To minimize aberrations, make sure that the refractive index of the mounting medium matches that of the immersion medium of the objective lens as much as possible.

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12. Images in 8-bit or 16-bit can be analyzed with the plug-in. Image deconvolution prior to running the analysis can improve their resolution and signal-to-noise ratio. 13. The X/Y spacing option in Multi Pinhole Search Option window makes sure that stray pixels belonging to the same pinhole are included. The default values for X/Y spacing option work quite well for nucleoid STED images too; however, it may be necessary to modify them depending on the image acquisition parameters. 14. The results window shows also the value related to the error of the fit, indicated as X (or Y) quality. As also suggested in the plug-in code documentation, values below 1 × 10-6 are considered satisfactory. 15. The pixel size can be easily retrieved from the metadata of the image dataset, or by checking the file properties in the imaging software. 16. The ROI can be of any shape. Although bulk analysis is possible with the plug-in, the software may freeze or shutdown if a rather large region is given.

Acknowledgments STED images were acquired at the CECAD imaging facility. E.M. is supported by the Deutsche Forschungsgemeinschaft (DFG, German Research Fundation)- SFB 1218 - grant number 269925409. References 1. Spinelli JB, Haigis MC (2018) The multifaceted contributions of mitochondria to cellular metabolism. Nat Cell Biol 20(7):745–754. https://doi.org/10.1038/s41556-0180124-1 2. Giorgi C, Marchi S, Pinton P (2018) The machineries, regulation and cellular functions of mitochondrial calcium. Nat Rev Mol Cell Biol 19(11):713–730. https://doi.org/10. 1038/s41580-018-0052-8 3. Scorrano L (2009) Opening the doors to cytochrome c: changes in mitochondrial shape and apoptosis. Int J Biochem Cell Biol 41(10): 1875–1883. https://doi.org/10.1016/j.bio cel.2009.04.016 4. Gustafsson CM, Falkenberg M, Larsson NG (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160. https://doi.org/10. 1146/annurev-biochem-060815-014402

5. Kauppila TES, Kauppila JHK, Larsson NG (2017) Mammalian mitochondria and aging: an update. Cell Metab 25(1):57–71. https:// doi.org/10.1016/j.cmet.2016.09.017 6. Kukat C, Davies KM, Wurm CA, Spahr H, Bonekamp NA, Kuhl I, Joos F, Polosa PL, Park CB, Posse V, Falkenberg M, Jakobs S, Kuhlbrandt W, Larsson NG (2015) Crossstrand binding of TFAM to a single mtDNA molecule forms the mitochondrial nucleoid. Proc Natl Acad Sci U S A 112(36): 11288–11293. https://doi.org/10.1073/ pnas.1512131112 7. Garrido N, Griparic L, Jokitalo E, Wartiovaara J, van der Bliek AM, Spelbrink JN (2003) Composition and dynamics of human mitochondrial nucleoids. Mol Biol Cell 14(4):1583–1596. https://doi.org/10. 1091/mbc.e02-07-0399 8. Kukat C, Wurm CA, Spahr H, Falkenberg M, Larsson NG, Jakobs S (2011) Super-resolution

Measurement of Nucleoid Size Using STED Microscopy microscopy reveals that mammalian mitochondrial nucleoids have a uniform size and frequently contain a single copy of mtDNA. Proc Natl Acad Sci U S A 108(33): 13534–13539. https://doi.org/10.1073/ pnas.1109263108 9. Hell SW, Wichmann J (1994) Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy. Opt Lett 19(11):780–782. https://doi.org/10.1364/ol.19.000780 10. Hell SW (2010) Far-field optical nanoscopy. In: Single molecule spectroscopy in chemistry, physics and biology. Springer, pp 365–398. https://doi.org/10.1007/978-3642-02597-6_19 11. Brown TA, Tkachuk AN, Shtengel G, Kopek BG, Bogenhagen DF, Hess HF, Clayton DA (2011) Superresolution fluorescence imaging of mitochondrial nucleoids reveals their spatial range, limits, and membrane interaction. Mol Cell Biol 31(24):4994–5010. https://doi. org/10.1128/MCB.05694-11 12. Stephan T, Roesch A, Riedel D, Jakobs S (2019) Live-cell STED nanoscopy of mitochondrial cristae. Sci Rep 9(1):12419. https://doi.org/10.1038/s41598-01948838-2 13. Silva Ramos E, Motori E, Bruser C, Kuhl I, Yeroslaviz A, Ruzzenente B, Kauppila JHK, Busch JD, Hultenby K, Habermann BH, Jakobs S, Larsson NG, Mourier A (2019) Mitochondrial fusion is required for regulation of mitochondrial DNA replication. PLoS

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Genet 15(6):e1008085. https://doi.org/10. 1371/journal.pgen.1008085 14. Nicholls TJ, Nadalutti CA, Motori E, Sommerville EW, Gorman GS, Basu S, Hoberg E, Turnbull DM, Chinnery PF, Larsson NG, Larsson E, Falkenberg M, Taylor RW, Griffith JD, Gustafsson CM (2018) Topoisomerase 3alpha is required for decatenation and segregation of human mtDNA. Mol Cell 69(1): 9–23 e26. https://doi.org/10.1016/j.molcel. 2017.11.033 15. Eggeling C, Hell SW (2015) STED fluorescence nanoscopy. In: Tinnefeld P, Eggeling C, Hell SW (eds) Far-field optical nanoscopy. Springer Berlin Heidelberg, Berlin, Heidelberg, pp 3–25 16. https://imagej.nih.gov/ij/index.html 17. https://imagej.nih.gov/ij/plugins/fwhm/ index.html 18. LeicaMicrosystems: the guide to STED sample preparation. https://www.leica-microsystems. com/science-lab/the-guide-to-sted-samplepreparation/ (2019) 19. Wurm CA, Neumann D, Schmidt R, Egner A, Jakobs S (2010) Sample preparation for STED microscopy. In: Papkovsky DB (ed) Live cell imaging: methods and protocols. Humana Press, Totowa, NJ, pp 185–199 20. Vicidomini G, Bianchini P, Diaspro A (2018) STED super-resolved microscopy. Nat Methods 15(3):173–182. https://doi.org/10. 1038/nmeth.4593

Part III Mitochondrial DNA Interacting Proteins

Chapter 10 How to Quantify DNA Compaction by TFAM with Acoustic Force Spectroscopy and Total Internal Reflection Fluorescence Microscopy Martial Martucci, Louis Debar, Siet van den Wildenberg, and Geraldine Farge Abstract Mitochondrial transcription factor A (TFAM) plays a key role in the organization and compaction of the mitochondrial genome. However, there are only a few simple and accessible methods available to observe and quantify TFAM-dependent DNA compaction. Acoustic Force Spectroscopy (AFS) is a straightforward single-molecule force spectroscopy technique. It allows one to track many individual protein-DNA complexes in parallel and to quantify their mechanical properties. Total internal reflection fluorescence (TIRF) microscopy is a high-throughput single-molecule technique that permits the real-time visualization of the dynamics of TFAM on DNA, parameters inaccessible with classical biochemistry tools. Here we describe, in detail, how to set up, perform, and analyze AFS and TIRF measurements to study DNA compaction by TFAM. Key words Acoustic Force Spectroscopy (AFS), Total Internal Reflection Microscopy (TIRF), mitochondrial DNA compaction, mitochondrial Transcription Factor A (TFAM), Single molecule biophysics

1

Introduction The mitochondrial genome is organized in protein-DNA complexes called nucleoids [1]. The most abundant protein found in these nucleoids is mitochondrial transcription factor A (TFAM) [2]. TFAM plays two crucial roles in mtDNA maintenance: (i) a role in transcription initiation, where it interacts specifically with the promotor regions of mtDNA [3]; and, (ii) a role in DNA compaction, for which it displays strong non-sequence-specific DNA binding [4, 5]. The latter function, DNA compaction, has been studied over the years using different techniques such as X-ray crystallography [6, 7], electron microscopy [8], or AFM [9].

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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With these techniques, researchers have established the structure of TFAM bound specifically and nonspecifically to DNA. They have also been able to propose different modes of compaction by TFAM, such as bending, loop formation, or bridging [10– 12]. However, these techniques do not typically explore the dynamics of the TFAM-DNA interactions. In contrast, single molecule techniques give the possibility to manipulate and follow DNA-TFAM filaments in real time, providing valuable tools to study their structure and dynamics. For instance, a combination of optical tweezers and fluorescence microscopy has been used to study the DNA compaction mode of WT and modified TFAM [13, 14]. It was also possible to show that TFAM diffuses rapidly on nonspecific DNA via a 1D sliding mechanism, which may provide a way for TFAM to locate the mtDNA promoters and initiate transcription [13]. Similarly, Tethered Particle Motion (TPM), a high-throughput single molecule technique, has successfully been used to study DNA compaction induced by WT TFAM and some TFAM variants [10, 13]. In this chapter, we describe two powerful, complementary, and high-throughput single molecule techniques to quantify and visualize TFAM-DNA interactions, namely AFS (Acoustic Force Spectroscopy) and TIRF (Total internal Reflection Fluorescence) microscopy. AFS is a newly developed technique that uses acoustic waves to apply force simultaneously on multiple single tethered molecules [15–18]. In our assay, we tether single DNA molecules between a glass slide and a bead. We first describe how to prepare a DNA substrate containing a portion of the mitochondrial genome and how to label it at one end of the DNA with digoxigenin and at the other end with biotin to attach it to the glass surface and to the bead, respectively. We then describe a typical AFS measurement to study DNA compaction. We first measure the RMS (Root mean square) values of the tethered molecules. The tethered beads undergo Brownian motion that is restricted by the reach of the DNA tether. Changes in the amplitude of these Brownian fluctuations will reflect a change in DNA tether length. By tracking the position of the tethered bead over time, we can determine the RMS and thus monitor the DNA length’s variation in real time. We then apply acoustic force to the tethers, and we perform an overstretching curve of DNA in order to determine the DNA’s persistence length (Lp). The persistence length of TFAM-coated DNA is considerably decreased compared to naked DNA molecules and thus constitutes a fingerprint of DNA compaction by TFAM [13]. In TIRF microscopy, an evanescent wave is created, which excites only the fluorophores near the glass surface. This results in the elimination of out-of-focus fluorescence, thus making it possible to achieve single molecule detection [19, 20]. TIRF microscopy allows the visualization of biological events at the single molecule scale, such as, for example, the real-time observation of rolling-

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circle DNA replication [21, 22]. In the second part of this chapter, we describe in detail how to perform a two-color TIRF experiment on DNA molecules coated with TFAM. We explain how to label a DNA template with biotins to attach it to the glass surface and with YOYO-1 to visualize it. We also provide a protocol to fluorescently label TFAM with Alexa-555. Finally, we describe step-by-step how to perform a TIRF experiment and how to analyze the obtained data.

2

Materials

2.1 TFAM Purification and Labeling

1. Purified recombinant human TFAM. 2. Dialysis buffer: 25 mM NaPO4 (pH 7), 10% glycerol, 0.2 M NaCl. 3. Dialysis device: Slide-A-Lyzer MINI dialysis units 7000 MWCO (Thermo Scientific). 4. Alexa Fluor 555 C2-maleimide (Invitrogen). 5. Illustra Microspin-G25 columns (GE Healthcare). 6. 1 M Dithiothreitol (DTT). 7. Spectrophotometer (NanoDrop or similar device).

2.2 DNA Production and Labeling

1. 10 μM Digoxygenin-labeled forward primer: 5′-[DIG]GCT [DIG]AAACCTAGCCCCAAACC-3′.

2.2.1 Template for AFS Experiments

2. 10 μM Biotin-labeled reverse primer: 5′- [Bio]T[BiodT] GTGTTGAGGGTTATGAGAGTAGC-3′. 3. LongAMP buffer (5X) (NEB) and LongAMP polymerase (NEB). 4. 10 mM dNTPs. 5. Nuclease-free water. 6. Template DNA (10–30 ng). 7. PCR clean-up Macherey-Nagel kit. 8. PCR tubes. 9. Thermocycler.

2.2.2 Template for TIRF Experiments

1. Klenow buffer (10X) and Klenow DNA polymerase exo(Promega). 2. 0.4 mM Biotin 14-dATP (Invitrogen). 3. 0.4 mM Biotin 14-dCTP (Invitrogen). 4. 10 mM dGTP and dTTP (Promega). 5. 250 μg/mL λDNA (Roche). 6. YOYO™-1 Iodide (491/509) (ThermoFisher).

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7. Nuclease-free water. 8. Illustra Microspin S-400 HR columns (GE Healthcare). 2.3 Preparation of the Flow Cells

1. Acetone.

2.3.1 For AFS Experiments

3. Alkaline hypochlorite solution: 0.2% sodium hypochlorite (Sigma).

2. 0.5 M Sodium thiosulfate.

4. PBS buffer: 137 mM NaCl, 2.7 mM KCl, 10 mM phosphate pH 7.4, supplemented with 5 mM NaN3 and 0.5 mM EDTA. 5. Anti-digoxygenin from sheep: 20 μg/mL in PBS (Roche). 6. 1% BSA (Sigma) in PBS buffer. 7. 5% Pluronic (F127) (Sigma) in PBS buffer. 8. 0.1% BSA and 0.25% Pluronic in PBS buffer. 9. 1% Casein (Sigma) in PBS buffer. 10. Buffer A: PBS buffer with 0.02% casein and 0.02% Pluronic. 11. Buffer B: 75 mM NaCl, 10 mM Tris–HCl pH 7.0, 0.5% glycerol. 12. Streptavidin-coated polystyrene beads (4.4 μm) (0.5%) (Spherotech). 13. 1 M NaOH. 14. Sigmacote. 15. Regal water: nitric acid and hydrochloric acid in a molar ratio of 1:3. 2.3.2

TIRF Experiments

1. Buffer C: 20 mM TrisOAc (pH 8), 20% sucrose, 10 mM dithiothreitol (DTT). 2. 10 mg/mL BSA-biotin (Sigma). 3. Alkaline hypochlorite solution: 0.2% sodium hypochlorite (Sigma). 4. 1 mg/mL Streptavidin (EMD Millipore). 5. 1.5 mg/mL Blocking reagent (Roche). 6. Nuclease-free water or MilliQ water. 7. 5 M NaCl. 8. 1 M NaOH. 9. Anhydrous DMSO.

2.4

Hardware

1. Isolation table. 2. Syringe pump (New Era Pump Systems). 3. AFS experimental setup: AFS experiments are performed on the AFS stand-alone G2 from LUMICKS B.V. It includes an inverted microscope, camera, controller box, and a stage

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Fig. 1 Simplified diagram of the experimental TIRF setup. Includes excitation lasers (Oxxius laser base), flow system (Lumicks), inverted microscope (IM) (Ti2E, Nikon), and detection system (Image splitter: Optosplit III, Cairn Research; EMCCD: iXon 888, ANDOR). Q: Quad Band TIRF filters (405/488/532/638–647), F1: filter 550/88, F2: filter 580/60, F3: filter 676/29, D1: dichroic 552 nm, D2: dichroic 650 nm

holding a chip holder and a chip. The AFS is connected to a desktop computer. 4. TIRF setup: TIRF experiments are performed on an inverted microscope TI2-E (Nikon) connected to a computer. The microscope contains the following parts (see schematic Fig. 1): • CFI Plan APO TIRF objective (100X, NA = 1.49, oil) (Nikon). • Excitation lasers: 405–100 mW, 488–100 532–100 mW, 638–150 mW (OXXIUS). • Highly sensitive camera for EMCCD (iXon 888, ANDOR).

fluorescence

mW,

detection:

• Image Splitter (Optosplit III, Cairn Research). • Excitation and emission filters optimized for the fluorophores of interest (Chroma). • Flow cell and flow cell holder (Lumicks). • Perfect focus system (PFS, Nikon). • 96-well plate holder (TI2-S-HW, Nikon). 2.5

Software

1. ImageJ. 2. MatLab. 3. NIS-Elements (Nikon). 4. Lumicks Tracking and Analysis software (Tracking software is freely available http://figshare.com/articles/AFS_soft ware/1195874).

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Methods

3.1 Assessing DNA Compaction by TFAM with AFS

The DNA template used is a ~9 kb fragment of the human mitochondrial genome (position 1671 to 10,600 nt) obtained and labeled by PCR. Amplification is performed using total DNA extracted from HEK cells (HEK 293, Invitrogen) and primers containing two biotins on one end and two digoxigenin on the other end (see Note 1).

3.1.1

Mix DNA template (10 ng of total cellular DNA), dNTPs (10 mM each final concentration), fwd and rev primers (10 μM each final concentration), LongAMP Taq polymerase (5 units), in 1X Long Amp PCR reaction buffer. Perform the amplification according to Table 1. Purify the PCR products with the Macherey-Nagel PCR cleanup kit. Elute DNA in 2 × 20 μL with elution buffer heated at 70 °C. Measure the DNA concentration on the Nanodrop and store at 4 ° C for a short period of time, or at -20 °C for longer term storage.

DNA Labeling

3.1.2 Flow Cell Functionalization

The flow cell used is a glass flow cell with one channel (volume ~ 5 ul). If handled gently, one flow cell can be reused for tens of experiments. The following cleaning and functionalization protocols are the manufacturer’s protocols with minor modifications and can be performed with the flow cell mounted on the microscope stage of AFS G2 setup (except the regal water step, which should be performed under a fume hood). For that, put the flow cell on the stage and connect the inlet to a reservoir containing the buffers and the outlet to the pump via a tubing (see Note 2). Use a flow rate of 100 μl/min for the washing, then, upon adding of the biological sample (from Subheading 3.1.2, step 2 onward), decrease the flow rate to 40 μL/min to avoid damaging the sample with too high shear/drag forces (see Note 3).

Table 1 PCR amplification for DNA labeling Temperature

Duration

Cycles

94 °C

30 s

1×

94 °C

15 s

30×

60 °C

30 s

65 °C

50 s/kb

65 °C

10 min

1×

10 °C

1

1×

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1. Cleaning: Connect a 3 mL syringe to the outlet of the flow cell. Insert 400 μL of alkaline hypochlorite solution. Incubate for 20 min (see Note 4). Insert 400 μL of 0.5 M sodium thiosulfate. Incubate for 5 min. Insert 400 μL of 1 M NaOH and incubate for 5 min. Rinse with nuclease-free water or MilliQ water between each step. Insert 50 μL of siliconizing reagent (Sigmacote) into the flow cell. Incubate for 1 min and air-dry the flow cell. Wash with acetone (see Note 5) and rinse thoroughly with nuclease-free water of MilliQ water. 2. Incubation of anti-digoxigenin: Insert 50 μL of antidigoxigenin (20 μg/mL) into the flow cell. Incubate for 20 min. 3. Passivation of the glass surface: Insert in the flow cell 250 μL of PBS solution with 0.1% BSA and 0.25% Pluronic. Incubate for 15 min. Repeat this step twice. 4. Anchoring DNA template to the glass surface: Insert 200 μL of buffer A in the flow cell. Then insert 50 μL of 0.02 ng/μL DNA solution in the flow cell and let it incubate for 20 min. Rinse with 200 μL of buffer A. 5. Washing the polystyrene beads: Add 10 μL of the bead solution to 1 mL of buffer A, mix by vortexing, and spin down the beads at 2000 × g for 2 min. Discard the supernatant. Resuspend the beads in 130 μL of buffer A. Repeat the washing step two times. 6. Attaching the beads: Gently insert the beads in the flow cell at a rate of 3 μL/min. Incubate 10 min. 3.1.3 AFS on TFAMCoated DNA

1. Getting started: Start the AFS and the tracking and analysis softwares (Lumicks). 2. Mounting the flow cell on the microscope stage: If this step has not been performed yet, first proceed as explained in Subheading 3.1.2. Then, when the flow cell is mounted, rinse the flow chamber with 400 μL buffer A to remove the beads that are not attached to DNA. Rinse the flow chamber with 50 μL buffer B. 3. Finding a field of view with tethered beads: Adjust the focus of the objective (20X) to a position where the beads appear out of focus (overfocus). Move the microscope stage to an area of the flow cell with multiple beads with a good spatial distribution (see Note 6). Select regions of interest (ROI) around the beads manually or automatically (see Note 7). Track the x and y positions of the beads using the Lumicks tracking software. For measuring the z position, make a look up table (LUT) (see Note 8). 4. Measuring RMS and FD curves on naked and TFAM-coated DNA. A typical experimental workflow is depicted in Fig. 2a.

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Fig. 2 AFS data analysis. (a) Schematic of the workflow of a typical AFS experiment. a: determination of the RMS for DNA tethers in the absence of TFAM and determination of the anchor point for the force calibration, b: force calibration, c: force ramp, d: incubation (with flow) with TFAM (this step can also be performed while applying a small force of about 2pN), e: determination of RMS for DNA tethers in the presence of TFAM, f: force ramp. (b) XY plot of tethered particles in the absence (black trace) and presence (gray trace) of 1 μM TFAM. The calculated RMS is 1.35 and 1.07 μm in the absence and presence of TFAM, respectively. (c) Histogram of

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Track the x and y positions of the beads (without force) for 15–30 min in the absence of protein. Typically, an x-y plot of a single tether is round (Fig. 2b) and the area probed by the tethered bead depends on the length of the tether. Calculate the anisotropic ratio α (Eq. 1) and the root mean square displacement (RMS) (Eq. 2) for all tethers using LUMICKS analysis software. α=

l major l minor

ð1Þ

where lmajor and lminor are the length of the major and minor axis of the x-y scatter plot, respectively [23]. sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi n h i 1X ð2Þ RMS = ðx - x Þ2 þ ðy - y Þ2 n i=1

where x and y are the coordinates of the bead position and n the total number of positions recorded [23]. Select tethers with an α below 1.4 (tethers with α above 1.4 are likely double tethers). For the selected tethers, determine the position where the DNA is anchored to the surface (i.e., the anchor point) from the bead position traces (see Note 9). Perform a force calibration by applying successively three different constant acoustic amplitudes to the DNA for 5 min each. For each amplitude, create a power spectrum of the bead position fluctuations and fit this power spectrum with quenched Brownian motion to obtain the force and subsequently deduce the Force/Amplitude ratio (use LUMICKS analysis software). After completing the force calibration, perform a force ramp (see Note 10) and generate a force-distance curve (Fig. 2d). Insert 1 μM TFAM (diluted in buffer B, see Note 11) in the flow cell at a flow rate of 0.6 μL/min for 15 min, stop the flow and then track again the x and y positions of the beads for 15–30 min. Determine the RMS for all tethers. Perform once again a force ramp and generate a force-distance curve for TFAM-coated DNA (see Note 12). At the end of the experiment, rinse the flow cell extensively with 1 M NaOH. Leave the NaOH overnight.

ä Fig. 2 (continued) the RMS values is obtained for one experiment; the values in the absence of TFAM are depicted in black and the values in the presence of TFAM are depicted in gray. (d) Typical force-extension curves for DNA in the absence (black trace) and presence (gray trace) of 1 μM TFAM. The pink dashed line represents the eWLC fit to our data, yielding a persistence length (Lp) of 43.0 nm in the absence of TFAM and 3.83 nm in the presence of TFAM and a contour length (Lc) of 2.78 in the absence of TFAM and 3.52 nm in the presence of TFAM, in agreement with earlier observations [13]. (e) DNA compaction by TFAM. TFAM molecules (gray) bind to DNA and compact it [5]

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5. Data representation and data analysis: To compare naked and TFAM-coated DNA molecules, plot the values of the average RMS with and without TFAM in a histogram (Fig. 2c). Fit the FD curves using the extensible wormlike chain model (eWLC) [24] which describes the dsDNA elastic behavior up to around 20 pN (Eq. 3) (we use a custom-written MatLab program). s ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi # "  F 1 kb T x = Lc 1 þ ð3Þ F Lp K: 2 where x is the extension (end-to-end distance) of the DNA, Lc is the contour length, kbT is Boltzmann’s constant times absolute temperature, F is the force, Lp the persistence length, and K the stretch modulus of DNA (set at 1200 pN). Determine the persistence length (Lp) and the contour length (Lc) of the DNA molecules in the presence and absence of TFAM. The decrease of the RMS and of the Lp both reflect DNA compaction upon TFAM binding (Fig. 2e). 3.2 Assessing TFAM Dynamics with TIRF Microscopy

In this protocol, we use λDNA, a linear molecule of 48,502 bp with two single-stranded 12-nt overhangs on both 5′-ends. λDNA is first labeled on both ends with biotins to attach it to the glass surface, then labeled with YOYO-1 for visualization. TFAM is labeled by covalently attaching a fluorophore to the -SH group (thiol) of the cysteines residues of the protein.

3.2.1 DNA Biotin Labeling

Mix λDNA (7 μg) with Klenow buffer (10X), Biotin 14-dATP (64 μM final concentration), Biotin 14-dCTP (64 μM final concentration), dGTP / dTTP (0.1 mM final concentration), and Klenow DNA polymerase exo- (5 units) in a total volume of 50 μL. Incubate 30 min at 37 °C. Then inactivate the reaction at 70 °C for 15 min. Purify the DNA on Illustra Microspin S-400 HR columns following the manufacturer’s protocol. Measure DNA concentration on the Nanodrop and store at 4 °C.

3.2.2

Dialyze TFAM in dialysis buffer for 1.5 h using a Slide-A-Lyzer MINI dialysis unit 7000 MWCO. Dissolve Alexa Fluor 555 C2-maleimide in anhydrous DMSO and add to the TFAM solution at a fivefold molar excess of dye over the protein. Incubate rotating at 4 °C for at least 2 h. The reaction should be protected from light as much as possible. Remove the excess dye using a G-25 Microspin column following the manufacturer’s protocol. Measure the degree of labeling spectrophotometrically (see Note 13). Add 1 mM DTT, aliquot the sample and store at -80 °C in aluminum foil after flash freezing in liquid nitrogen.

TFAM Labeling

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The flow cell holder is mounted on a 96-well plate holder. The inlet of the flow cell is connected to a 3 mL syringe and the outlet of the flow cell is connected to a pump. The cleaning and DNA anchoring protocols are adapted from [25]. 1. Cleaning: All the cleaning steps are performed at a flow rate of 50 μL/min. Wash the flow cell with alkaline hypochlorite solution for 20 min (see Note 14). Rinse with nuclease-free water. Insert 200 μL of 1 M NaOH and incubate 10 min. Rinse with nuclease-free water. Wash with 200 μL of Buffer C. 2. Flow cell passivation: Insert 250 μL of Buffer C supplemented with 1 mg/mL BSA-Biotin in the flow cell and incubate for 15 min. Repeat this step twice. Insert 250 μL of Buffer C supplemented with 0.1 mg/mL streptavidin in the flow cell and incubate 15 min. Repeat this step twice. Insert 300 μL of blocking reagent in the flow cell and incubate for 15 min. Rinse the flow cell with 200 μL buffer C.

3.2.4 Anchoring DNA Template to the Glass Surface

Mix biotinylated λDNA (0.25 ng/μL final concentration) with YOYO-1 (0.5 nM) in buffer C. Insert 100 μL of this solution in the flow cell (40 μL/min) and incubate for 20 min. Flush in at least 200 μL of buffer C at 100 μL/min to stretch the DNA molecules at the surface of the flow cell (see Note 15). Rinse with buffer C to remove the YOYO-1 in excess.

3.2.5 Sample Preparation

Dilute labeled TFAM in buffer C without salt to a concentration of 5–20 nM (see Notes 16 and 17). Insert 100 μL of diluted labeled TFAM in the flow cell (50 μL/min) and incubate for 5 min. Wash with buffer C supplemented with 25 mM NaCl to remove the excess of free TFAM. Start imaging immediately.

3.2.6

Imaging

Images are acquired in total internal reflection mode, through a Nikon Plan APO TIRF objective (see Subheading 2.4), on an EMCCD camera. The images are split using an image splitter into separate spectral components which are aligned side by side on the camera chip (size of each image is 1024 × 340 pixels) (Fig. 3a). Start recording a movie. Excite the YOYO-1 with the 488 nm laser (laser power: 9 mW, exposure time: 500 ms, gain 200, 2–10 frames) (see Note 18). Turn off the 488 nm laser and turn on the 532 nm laser to excite Alexa555 (laser power: 2.6 mW, exposure time: 500 ms, gain 200, 40–300 frames). Record until the molecules of interest are bleached (see Note 19). Stop recording the movie. Move to another area of the flow cell and record another movie.

3.2.7

Data Analysis

Quantifying the Intensity of a Single Fluorophore Take a DNA molecule with a dense TFAM coverage (see Note 17). For each visible fluorescent TFAM particle, plot the intensity of the particle versus time. Use bleaching steps to determine the intensity of a

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Fig. 3 TIRF data analysis. (a) Two color TIRF fluorescence images of a DNA molecule coated with TFAM. Upper image (green): DNA labeled with YOYO-1 excited with the 488 nm laser. Middle image (blue): Fluorescent TFAM-Alexa555 excited with 532 nm laser. Lower image: Merge of the two images. Scale bar: 1 μm. (b) Intensity histogram of single step bleaching. The fluorescence intensity was measured for 103 fluorescent particles. We obtain a mean fluorescence value for a single fluorophore of approximately 2000 a.u. (c) Kymograph created from TFAM spots on a DNA molecule (as shown in (a)) showing the diffusion of fluorescent TFAM on DNA. Time (s) and distance (μm) are indicated at the bottom and left of the kymograph, respectively. (d) Representative trajectory generated from tracking the motion of a single TFAM on DNA. (e) Mean-squared displacement (MSD) of a TFAM molecule (same molecule as in (d)) versus time interval (τ). The diffusion coefficient calculated for this molecule from the linear fit (red line) to the MSD plot is 7 × 103 nm2 s-1

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single fluorophore. Perform this analysis on at least 30 molecules. Build a histogram of the steps found (example Fig. 3b) (see Note 20). Creating a Kymograph Select a DNA molecule with a sparse TFAM coverage. Create a kymograph (see Note 21 and Fig. 3c) for each DNA molecule selected. Use the tool “Image>ND processing>create a kymograph by line” if using the Nikon software (see Note 22). Selecting Single TFAM Molecules Determine the initial intensity of the fluorescent particles detectable on the kymograph. Select the molecules with an intensity corresponding to a single TFAM (i.e., to two fluorophores, see Note 13). Tracking Single Particles Track single TFAM molecules (see Note 23). Take only traces with a minimal length of 30 frames. Determine the trajectory of the moving spots (Fig. 3d), and the MSD (Fig. 3e). Calculate the diffusion constant (D) from the MSD plot (MSD = 2Dτ + offset), where τ is the time interval and the offset indicates the position accuracy.

4

Notes 1. We chose a portion of mtDNA that does not contain any specific binding sites, but other regions of mtDNA can be amplified following the same protocol and using primers flanking the region of interest. For PCR amplification of a portion of human mtDNA, it is possible to use as template DNA extracted from any other human cell line available in the lab. 2. Make sure to always keep liquid in the tank. To improve the flow in the tubing, wash the microtube with ethanol before connecting it to the flow cell outlet. 3. The washing steps can be performed by directly connecting a syringe to the outlet of the flow cell. The pump can be connected only at step 2 of Subheading 3.1.2. 4. In case a bacterial lawn remains stuck to the surface, insert 500 μL of 25% regal water in the flow cell and incubate for 15 min. 5. Be careful to not drop acetone on the piezo element as it can damage welding spots. 6. The beads should not be too close, otherwise they will interfere with each other in the tracking, yet there should be enough beads in the field of view, typically in the order of 30 beads.

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7. The diffraction rings of the beads should be included in the ROI. The size of the ROI will depend on many parameters, such as the bead size, the objective used, etc. Typically for a 4.4 μm bead and using a 20× magnification objective, we select ROIs of about 60 nm. For the automatic bead detection, a reference bead has to be preselected first. 8. The range of the LUT has to be bigger than the size of the extended DNA molecule. In our case, the DNA molecule is 3 μm long and we make the LUT over 10 μm. 9. The anchor point should be carefully determined for a good force calibration. 10. For the force ramp, we usually set a linear force ramp (back and forth) of 5000 steps, over 30 s, and the maximum force reached at 15 pN. 11. Buffer B contains glycerol as TFAM stock protein is stored in 10% glycerol. Be careful not to use more than 0.5% glycerol in the assay, as a higher glycerol concentration may result in more surface interactions. In case the protein is stored in high glycerol concentration, or the volume of protein used in the assay is important, first dialyze the protein against a buffer containing not more than 0.5% glycerol. 12. To clearly observe the effect of the protein on the DNA, make sure to measure the RMS and the force-distance on the same good tethers before and after addition of TFAM. 13. Wild-type human TFAM contains two cysteines. We typically obtain a ratio of two dyes per TFAM monomer, suggesting that the two cysteines of TFAM are exposed to the solvent and can be labeled. 14. In case of bacterial contamination, this step can be performed overnight. It is also possible to use regal water diluted in water (1:3 v/v) for 15 min. 15. If the DNA is not stretched enough, it is possible to stop the flow 2 or 3 times for 2 min during this step. 16. Always keep TFAM and the TFAM dilutions on ice to prevent loss of function. Perform the dilution of TFAM just prior to use and briefly centrifuge the sample to remove possible aggregates. 17. To obtain single-molecule resolution and to be able to follow the diffusion of the TFAM molecules on DNA, the concentration of TFAM used should be low, resulting in a sparse coverage of the DNA (typically we use 5 nM). On the contrary, for measuring the intensity of a single fluorophore, it is better to start with a DNA molecule with a denser TFAM coverage, so we use a higher TFAM concentration (typically we use 20 nM). At these concentrations, TFAM forms patches that are

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immobile, making it possible to follow its fluorescence in time without the risk that it is blurred by any movement [13]. 18. This is just to determine the position of the DNA molecules, so only take a few frames. 19. The laser power and exposure time should be chosen according to the process that is investigated. For example, to obtain a diffusion coefficient for molecules moving along the DNA molecule, the movies should be significantly long (in our experience at least 30 frames) and laser power should thus be low to limit photobleaching. In contrast, fast processes such as binding/unbinding may require short exposure times (and consequently higher laser powers). 20. In our hands a single fluorophore has an intensity of about 2000 a.u. 21. A standard way of following a particle moving in one dimension, in this case to follow the diffusion of TFAM molecules on DNA, is to create a kymograph. A kymograph takes the pixels along a DNA molecule in the video and stacks them as time progresses. 22. There are also many free plug-ins available for ImageJ to create kymographs. 23. There are many different tracking softwares available in the single molecule labs or otherwise in the literature. Some tracking softwares are based on 2D Gaussian fit of the fluorescent spot, others on applying image filtering (e.g., the ImageJ plugin “Particle Tracker 2D/3D” [26]).

Acknowledgments The authors thank Maria Falkenberg for valuable support. This work was supported by the University Clermont Auvergne (UCA), the Laboratoire de Physique de Clermont (LPC), the Region Auvergne-Rhoˆne-Alpes et l’Union Europe´enne dans le cadre du Fonds Europe´en de De´veloppement Re´gional (FEDER), and the Association Franc¸aise Contre les Myopathies Te´le´thon (AFM) (#21411 to GF and #22765 to LD). References 1. Spelbrink JN (2010) Functional organization of mammalian mitochondrial DNA in nucleoids: history, recent developments, and future challenges. IUBMB Life 62:19–32. https://doi.org/10.1002/iub.282 2. Gustafsson CM, Falkenberg M, Larsson N-G (2016) Maintenance and expression of

mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160. https://doi.org/10. 1146/annurev-biochem-060815-014402 3. Falkenberg M, Gaspari M, Rantanen A, Trifunovic A, Larsson N-G, Gustafsson CM (2002) 2 Mitochondrial transcription factors B1 and B2 activate transcription of human

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mtDNA. Nat Genet 31:289–294. https://doi. org/10.1038/ng909 4. Wang YE, Marinov GK, Wold BJ, Chan DC (2013) Genome-wide analysis reveals coating of the mitochondrial genome by TFAM. PLoS One 8:e74513. https://doi.org/10.1371/ journal.pone.0074513 5. Farge G, Falkenberg M (2019) Organization of DNA in mammalian mitochondria. Int J Mol Sci 20. https://doi.org/10.3390/ ijms20112770 6. Ngo HB, Kaiser JT, Chan DC (2011) The mitochondrial transcription and packaging factor Tfam imposes a U-turn on mitochondrial DNA. Nat Struct Mol Biol 18:1290–1296. https://doi.org/10.1038/nsmb.2159 7. Rubio-Cosials A, Sidow JF, Jime´nezMene´ndez N, Ferna´ndez-Milla´n P, Montoya J, Jacobs HT, Coll M, Bernado´ P, Sola` M (2011) Human mitochondrial transcription factor A induces a U-turn structure in the light strand promoter. Nat Struct Mol Biol 18:1281–1289. https://doi.org/10. 1038/nsmb.2160 8. Kukat C, Davies KM, Wurm CA, Spa˚hr H, Bonekamp NA, Ku¨hl I, Joos F, Polosa PL, Park CB, Posse V, Falkenberg M, Jakobs S, Ku¨hlbrandt W, Larsson N-G (2015) Crossstrand binding of TFAM to a single mtDNA molecule forms the mitochondrial nucleoid. Proc Natl Acad Sci U S A 112:11288–11293. https://doi.org/10.1073/pnas.1512131112 9. Kaufman BA, Durisic N, Mativetsky JM, Costantino S, Hancock MA, Grutter P, Shoubridge EA (2007) The mitochondrial transcription factor TFAM coordinates the assembly of multiple DNA molecules into nucleoid-like structures. Mol Biol Cell 18: 3225–3236. https://doi.org/10.1091/mbc. e07-05-0404 10. Ngo HB, Lovely GA, Phillips R, Chan DC (2014) Distinct structural features of TFAM drive mitochondrial DNA packaging versus transcriptional activation. Nat Commun 5: 3 0 7 7 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncomms4077 11. Rubio-Cosials A, Battistini F, Gansen A, Cuppari A, Bernado´ P, Orozco M, Langowski J, To´th K, Sola` M (2018) 9 protein flexibility and synergy of HMG domains underlie U-turn bending of DNA by TFAM in solution. Biophys J 114:2386–2396. https://doi. org/10.1016/j.bpj.2017.11.3743 12. Malarkey CS, Bestwick M, Kuhlwilm JE, Shadel GS, Churchill MEA (2012) Transcriptional activation by mitochondrial transcription factor A involves preferential distortion of promoter

DNA. Nucleic Acids Res 40:614–624. https:// doi.org/10.1093/nar/gkr787 13. Farge G, Laurens N, Broekmans OD, van den Wildenberg SMJL, Dekker LCM, Gaspari M, Gustafsson CM, Peterman EJG, Falkenberg M, Wuite GJL (2012) Protein sliding and DNA denaturation are essential for DNA organization by human mitochondrial transcription factor A. Nat Commun 3:1013. https://doi.org/ 10.1038/ncomms2001 14. King GA, Hashemi Shabestari M, Taris K-KH, Pandey AK, Venkatesh S, Thilagavathi J, Singh K, Krishna Koppisetti R, Temiakov D, Roos WH, Suzuki CK, Wuite GJL (2018) Acetylation and phosphorylation of human TFAM regulate TFAM-DNA interactions via contrasting mechanisms. Nucleic Acids Res 46:3633–3642. https://doi.org/10.1093/ nar/gky204 15. Kamsma D, Creyghton R, Sitters G, Wuite GJL, Peterman EJG (2016) Tuning the music: acoustic force spectroscopy (AFS) 2.0. Methods 105:26–33. https://doi.org/10. 1016/j.ymeth.2016.05.002 16. Sitters G, Kamsma D, Thalhammer G, RitschMarte M, Peterman EJG, Wuite GJL (2015) Acoustic force spectroscopy. Nat Methods 12: 47–50. https://doi.org/10.1038/nmeth. 3183 17. Kamsma D, Wuite GJL (2018) Singlemolecule measurements using acoustic force spectroscopy (AFS). Methods Mol Biol 1665: 341–351. https://doi.org/10.1007/978-14939-7271-5_18 18. Lin S-N, Qin L, Wuite GJL, Dame RT (2018) Unraveling the biophysical properties of chromatin proteins and DNA using acoustic force spectroscopy. Methods Mol Biol 1837:301– 316. https://doi.org/10.1007/978-1-49398675-0_16 19. Kudalkar EM, Davis TN, Asbury CL (2016) Single-molecule total internal reflection fluorescence microscopy. Cold Spring Harb Protoc 2016. https://d oi.org/10.1101/pdb . top077800 20. van den Wildenberg SMJL, Prevo B, Peterman EJG (2018) A brief introduction to singlemolecule fluorescence methods. Methods Mol Biol 1665:93–113. https://doi.org/10.1007/ 978-1-4939-7271-5_5 21. Tanner NA, van Oijen AM (2009) Singlemolecule observation of prokaryotic DNA replication. Methods Mol Biol 521:397–410. https://doi.org/10.1007/978-1-60327-8157_22 22. Tigano M, Phillips AF, Sfeir A (2020) Singlemolecule analysis of mtDNA replication with

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Chapter 11 Assessing TFAM Binding to Human Mitochondrial DNA Takehiro Yasukawa and Dongchon Kang Abstract Mitochondrial transcription factor A (TFAM) is a mitochondrial DNA (mtDNA)-binding protein that plays a crucial dual role in the initiation of mitochondrial transcription initiation and mtDNA maintenance. Because TFAM directly interacts with mtDNA, assessing its DNA-binding property can provide useful information. This chapter describes two in vitro assay methods, an electrophoretic mobility shift assay (EMSA) and a DNA-unwinding assay with recombinant TFAM proteins, which both require simple agarose gel electrophoresis. These are used to investigate the effects of mutations, truncation, and posttranslational modifications on this key mtDNA regulatory protein. Key words Mitochondrial transcription factor A, TFAM, Mitochondrial DNA, mtDNA, Mitochondrial nucleoids, Electrophoretic mobility shift assay, EMSA, DNA-unwinding assay

1

Introduction It is generally accepted that mitochondrial transcription factor A (TFAM) is a dual-function protein, as both a mitochondrial transcription initiation factor and the major mitochondrial nucleoid protein. Inside mitochondria, TFAM initiates transcription initiation through binding to specific positions on mitochondrial DNA (mtDNA), upstream of heavy- and light-strand promotors (HSP and LSP) [1, 2]. This is followed by assembly of mitochondrial RNA polymerase (POLRMT/mtRNAP) and the second mitochondrial transcription initiation factor, mitochondrial transcription factor B2 (TFB2M/mtTFB2), to form ternary mitochondrial transcription initiation complexes at the transcription start sites [3]. TFAM has a second function in which it binds in a sequencenonspecific manner to the entire mtDNA molecule [4, 5], providing stability to the multicopy mitochondrial genome [6]. Consistently, TFAM expression levels are high enough to fulfil this function [4, 7]. Crystal structural analysis shows that doublestranded DNA oligonucleotides with an HSP or LSP sequence and a nonspecific sequence have a similar 180° U-turn conformation

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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when bound by TFAM [8]. Although TFAM acts as an architectural component of mitochondrial nucleoids through sequencenonspecific binding, how the same protein plays a role in the initiation of transcription at the HSP and LSP specifically is unclear. In vitro assays that can investigate the DNA-binding of TFAM may be an appropriate tool to answer this question. In addition, because in vitro assays can be performed using recombinantly expressed proteins, researchers can introduce mutations [9], protein domain deletion [10], and posttranslational modifications [11] to TFAM to evaluate their effects in the assays. When studying the DNA-binding properties of TFAM, two elements need to be considered: the DNA-binding activity and ability to change the DNA topology, which are involved in the introduction of negative supercoiling to mtDNA (see the Discussion section in [11] and the references therein), and compaction of mtDNA in mitochondrial nucleoids (see Chapter 10). This chapter describes the experimental procedures of an electrophoretic mobility shift assay (EMSA) and a DNA-unwinding assay, using recombinant human TFAM proteins and 3 kb plasmid DNA as a downsized model of circular human mtDNA. We recently used these to investigate the effects of lysine acetylation on TFAM [11]. There are more sophisticated in vitro assays [9, 12]. Quantitative kinetic analysis of TFAM binding to specific mtDNA sequences can be performed using a surface plasmon resonance instrument [10, 13]. Chromatin immunoprecipitation methods have been applied to mtDNA to assess the overall TFAM binding in living cells [5, 10, 13]. The two assays described here are en masse experimental settings that enable researchers to study the global TFAM–DNA interaction in vitro. Moreover, these assays do not require special instruments, but can be conducted using a standard agarose gel electrophoresis system.

2

Materials

2.1 General Consideration

2.2 Agarose Gel and Running Buffer

All solutions should be prepared using ultrapure water (Milli-Q water) and analytical-grade reagents free from DNase. In addition, the electrophoresis apparatus and glassware for solution preparation should be DNase free, because even a single bit of DNA nicking of the plasmid DNA will eliminate the topological information about the plasmid. Wear a mask and handle samples and apparatus with clean hands and clean gloves to avoid introducing nucleases in the experimental system. 1. 1× TAE buffer: 40 mM Tris–HCl, 40 mM acetic acid, and 1 mM ethylenediaminetetraacetic acid (EDTA), pH 8.0. 2. 50× TAE buffer: 2 M Tris–HCl, 2 M acetic acid, and 50 mM EDTA, pH 8.0.

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3. Gel running buffer: 1× TAE buffer with ethidium bromide (EtBr) added where necessary at a final concentration of 0.5 μg/mL. 4. 1% agarose gel with 1× TAE for EMSA and DNA-unwinding assay. 5. 1% agarose gel with 1× TAE and EtBr at a final concentration of 0.5 μg/mL for Southern hybridization. 6. Erlenmeyer flask. 7. Microwave. 8. 6 cm long and 10.5 cm wide gel trays with a gel comb. 9. 10 mg/mL EtBr. 2.3 Preparation of Proteins and Plasmid DNA

1. TFAM: mature human TFAM-mimicking protein with the histidine tag-containing polypeptide at the N-terminus (see Note 1). The expression plasmid construct is a DNA fragment containing complementary DNA (cDNA) of 44–246 amino acid residues of human TFAM, cloned between BamHI and EcoRI sites of pPRO-EX-HTb vector (Invitrogen) [11]. 2. Plasmid DNA: pBluescript KS(-) (3 kb), but other plasmids can be used as well. 3. Calf-thymus-derived DNA topoisomerase I (TaKaRa Bio Inc.). 4. Proteinase K: a DNase-free grade enzyme must be used. 5. Commercially available plasmid preparation kit (see Note 5). 6. Restriction enzyme (EcoRV). 7. Phenol (saturated with TE buffer). 8. Chloroform-isoamyl alcohol (24:1). 9. 100% and 70% ethanol (EtOH). 10. 1× phosphate-buffered saline (PBS) for 1× PBS/20% glycerol. 11. 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-NaOH (pH 7.2). 12. 3 M NaOAc (pH 5.2). 13. Tris–HCl. 14. KCl. 15. MgCl2. 16. Dithiothreitol (DTT). 17. Spermidine. 18. Bovine serum albumin (BSA). 19. EDTA. 20. HisTrap FF crude, HiTrap Heparin HP, and HiLoad 16/600 ¨ KTAprime plus. Superdex 75 pg columns using A 21. Centrifuge.

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2.4 DNA Detection After Electrophoresis

1. Gel staining solution: Sybr Green I solution diluted with 1× TAE 10,000 times. Prepare this solution freshly during agarose gel electrophoresis, preferably just before stopping the run. 2. Imaging analyzer equipped with an appropriate-wavelength laser and filter to detect Sybr Green I, for example, an ImageQuant LAS4000 instrument. 3. Depurination buffer: 0.25 N HCl. 4. Denaturing buffer: 1.5 M NaCl, 0.5 M NaOH. 5. Neutralizing buffer: 0.5 M Tris-HCl pH 7.4, 1.5 M NaCl (see Note 2). 6. Buffers for Southern hybridization are described in the manufacturer’s instructions for the kit used. 7. Nylon membrane (Hybond N+). 8. Hybridization probe. 9. A region of pBluescript KS(-) plasmid can be PCR-amplified for detection with a primer pair (5′-GAGCGCAGAAGTGGTCCTG-3′ and 5′-ACATTTCCGTGTCGCCCTTATTC-3′) [11]. The PCR product should be agarose-gel-purified.

3

Methods

3.1 Preparation of Agarose Gel Running Buffer

1. Prepare concentrated 50× TAE buffer and dilute with Milli-Q water to prepare 1× TAE buffer. Store both 50× and 1× TAE buffers at room temperature. For EtBr-containing gel electrophoresis, add EtBr to 1× TAE buffer at a final concentration of 0.5 μg/mL.

3.2 Preparation of Agarose Gel

1. Weigh 1% (w/v) agarose powder and mix with 1× TAE buffer in an Erlenmeyer flask (e.g., 1 g of agarose powder with 100 mL of 1× TAE buffer). 2. Dissolve the agarose completely using a microwave. 3. If making an EtBr-containing gel, add EtBr to the gel solution at a final concentration of 0.5 μg/mL and mix. 4. Set the gel solution in a gel tray with a gel comb (see Note 3).

3.3 Preparation of Recombinant TFAM

1. Express recombinant TFAM in Escherichia coli (E. coli) BL21 cells by isopropyl β-d-1-thiogalactopyranoside (IPTG) induction. 2. Sonicate the cells and recover the soluble fraction. 3. Purify TFAM from the soluble fraction. Perform sequential column chromatography with HisTrap FF crude, HiTrap

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Heparin HP, and HiLoad 16/600 Superdex 75 pg columns ¨ KTAprime plus to obtain a highly pure TFAM using A preparation. 4. Determine the concentration of the purified TFAM (see Note 4). 5. Store the purified TFAM in 1× PBS/20% glycerol in small aliquots at -80 °C [11]. 3.4 Preparation of Plasmid DNA

1. Prepare plasmid DNA using competent E. coli cells, such as DH5α, and a commercially available plasmid preparation kit (see Note 5).

3.5 Electrophoretic Mobility Shift Assay (EMSA)

To evaluate the DNA-binding affinity of TFAM, perform EMSA using linearized plasmid DNA to minimize the effect of DNA topological constraints on TFAM binding (see Note 6) [11]. Figure 1 shows how the extent of retardation of the fragments is examined. 1. Incubate plasmid DNA (see Note 7) at 37 °C for 1 h with a restriction enzyme (EcoRV) to cut the plasmid once. 2. Add an equal volume of phenol (saturated with TE buffer) to the reaction mixture. (A)

linear DNA

TFAM

AGE & Sybr Green I staining

(B) 8 7 6 5 4 3 2 1.65 (kb)

M U Lin (−) 56.5 22.6 11.3 7.5 5.6 4.5 3.8 3.2 2.8

3.5.1 Preparation of Linearized Plasmid

Fig. 1 Experimental scheme and an example result of electrophoretic mobility shift assay (EMSA). (a) Schematic drawing of EMSA procedure. Double-stranded DNA is depicted as a single line. Agarose gel electrophoresis is abbreviated to AGE. (b) An EMSA result with TFAM using 3 kb linearized DNA fragments. M: DNA ladder maker, U: uncut plasmids, Lin: linearized plasmids, (-): mock incubation (no TFAM was included in the reaction). Numbers below lanes represent the molecular ratios of TFAM to DNA fragments: 1 TFAM/56.5, 22.6, 11.3, 7.5, 5.6, 4.5, 3.8, 3.2, and 2.8 bp of the plasmid DNA. (The gel image is a reproduction of Fig. 1a of reference [11])

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3. Shake the tube vigorously by hand for 3 min and centrifuge at 17,900 × g for 3 min at room temperature. 4. Recover the water phase to a fresh microtube, and add an equal volume of chloroform-isoamyl alcohol (24:1) to it. 5. Shake the tube vigorously by hand for 3 min, and centrifuge the mixture at 17,900 × g for 3 min at 4 °C. Recover the water phase to a fresh microtube (see Note 8). 6. Add 1/10 (v/v) 3 M NaOAc (pH 5.2) to the recovered water phase and mix well, then add 2.5 (v/v) 100% EtOH (precooled at -20 °C to -25 °C) and mix well again (see Note 9). Incubate the mixture at -20 °C to -25 °C for at least 1 h. 7. Centrifuge the microtube at 17,900 × g for 20 min at 4 °C, before removing the supernatant carefully so as not to disturb the DNA pellet. 8. Add 70% EtOH (precooled) gently and invert the tube a couple of times gently. 9. Centrifuge the tube at 17,900 × g for 5 min at 4 °C, before removing the supernatant carefully and completely. 10. Air-dry the pellet for ~25 min (see Note 10). 11. Finally, dissolve the pellet with 10 mM HEPES-NaOH (pH 7.2) and measure the DNA concentration using a UV spectrophotometer or other measurement method. 3.5.2 TFAM-Binding Incubation

1. Make a 20 μL reaction mixture containing 35 mM Tris-HCl pH 8.0, 72 mM KCl, 5 mM MgCl2, 5 mM DTT, 5 mM spermidine, 0.1 mg/mL of BSA, and 10 mM EDTA (see Note 11). 2. Incubate different amounts of TFAM proteins with 125.5 ng of linearized plasmid in the reaction mixture at 37 °C for 30 min. The TFAM amount will vary depending on the purpose of the experiment and on the introduction of a mutation, depletion, or modification in the protein. Adding 0, 100, 250, 500, 750, 1000, 1250, 1500, 1750, and 2000 ng of TFAM to the reaction mixtures obtains ratios of 1 TFAM per 56.5, 22.6, 11.3, 7.5, 5.6, 4.5, 3.8, 3.2, and 2.8 bp, respectively, of the linearized plasmid DNA [11].

3.5.3 Agarose Gel Electrophoresis and Detection of TFAM-Bound DNA

1. Pour 1× TAE gel running buffer at room temperature into the electrophoresis apparatus and submerge the agarose gel (see Note 12). Neither buffer nor gel should contain EtBr. 2. Aliquot 8 μL of the 20 μL reaction mixture from Subheading 3.5.2 and add 0.89 μL of 10× loading buffer. 3. Load the samples to the gel and electrophorese at 66 mA current for ~4.5 h at room temperature (see Note 13).

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4. After electrophoresis, transfer the gel from the electrophoresis tank to a container containing the gel staining solution, and gently rock the gel in the solution for 30 min (see Note 14). 5. Detect the bands using an imaging analyzer equipped with an appropriate-wavelength laser and filter to detect Sybr Green I, for example, an ImageQuant LAS4000 instrument. 3.6 DNA-Unwinding Assay

(A)

To study the DNA-unwinding activity of TFAM (the ability to change the DNA topology toward negative supercoiling, or toward reducing DNA’s double-helical turn), perform the DNA-unwinding assay using a closed circular plasmid and eukaryotic DNA topoisomerase I (see Note 15) [10, 11]. The topological status of the plasmid DNA is evaluated with electrophoretic migration in an agarose gel (Fig. 2a, b). For accurate evaluation of TFAM

Plasmid DNA

Supercoiled

Topoisomerase I

Proteinase K treatment

* Relaxed

*

AGE & Sybr Green I staining (B) Supercoiled

AGE(+EtBr) & Southern hybridization (C)

Relaxed

(B)

(C) OC NC

L

L

SC

Lin U (−) 56.5 22.6 11.3 7.5 5.6 4.5 3.8 3.2 2.8

SC

M Lin U (−) 56.5 22.6 11.3 7.5 5.6 4.5 3.8 3.2 2.8

4 3 2 1.65 (kb)

Fig. 2 Experimental scheme and example results of DNA-unwinding assay. (a) Schematic drawing of the DNA-unwinding assay procedure. Asterisks indicate the continuation of the procedure. (b, c) Example results of DNA-unwinding assay and Southern hybridization. Lane labeling below the gel images is the same as in Fig. 1. L linear form, SC (fully) supercoiled, OC open circular (the relaxed form with no nick), NC nicked circular. (The images in b and c are reproductions of Fig. 1c and Supplementary Fig. 1b of reference [11], respectively)

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DNA-unwinding assay results, it is necessary to confirm that no DNA nicking of the plasmid DNA occurs in the unwinding assay by running the DNA in an EtBr-containing agarose gel (see Note 16). 3.6.1

Unwinding Assay

For the DNA-unwinding assay, use untreated (uncut) plasmid DNA. 1. Make a 20 μL reaction mixture comprising 35 mM Tris-HCl pH 8.0, 72 mM KCl, 5 mM MgCl2, 5 mM DTT, 5 mM spermidine, 0.1 mg/mL of BSA, and 10 mM EDTA (Note 11). 2. Incubate 125.5 ng of the plasmid DNA preparation with 0.5 units of calf thymus DNA topoisomerase I (see Note 17) in the reaction mixture at 37 °C for 30 min. 3. Add different amounts of TFAM (0–2000 ng) to the reaction mixtures containing plasmid DNA and topoisomerase I, and incubate at 37 °C for another 30 min. The TFAM amount range should be determined as discussed in Subheading 3.5.2. 4. Add proteinase K at a final concentration of 167 μg/mL to the reaction mixtures and incubate them at 37 °C for 20 min. 5. Then, add an equal volume of chloroform-isoamyl alcohol (24: 1), vigorously shake the tubes by hand for 3 min, and centrifuge them at 17,900 × g for 3 min at 4 °C. Recover the water phase containing the plasmid DNA; this step removes proteins from plasmid DNA.

3.6.2 Agarose Gel Electrophoresis and Plasmid DNA Visualization

Procedure is essentially the same as in Subheading 3.5.3. 1. Aliquot appropriate amounts of the samples prepared in Subheading 3.6.1 and mix with 10× loading dye. 2. Perform electrophoresis at room temperature. Neither buffer nor gel should contain EtBr. Example running conditions are 69 mA constant current for 3.5 h, but optimization with specific electrophoresis apparatus may be necessary (see Note 13).

3.6.3 Confirmation of the Absence of DNA Nick Introduction by Southern Hybridization

1. Prepare 1% agarose gel and 1× TAE gel running buffer, both containing 0.5 μg/mL of EtBr. 2. Run the plasmid samples subjected to the DNA-unwinding assay at room temperature. 3. After electrophoresis, transfer the gel into a glass container with depurination buffer and rock for 20 min. 4. Then, rock the gel twice in denaturing buffer for 15 min and twice in neutralizing buffer for 15 min. 5. Blot the gel to a nylon membrane with a standard Southern blotting procedure overnight.

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6. The following day, air-dry the membrane and fix the DNA on it (see Note 18). 7. Perform Southern hybridization to detect the plasmid DNA using a hybridization probe (see Note 19).

4

Notes 1. With regard to recombinant TFAM expression, make sure that the mitochondrial targeting signal (MTS) is not included in the expressed protein. Native TFAM is translated in the cytoplasm of human cells in premature form with the MTS at the N-terminal. The premature form is imported into the mitochondria, where the MTS is cleaved, and TFAM changes into its mature form. In human TFAM, the first 42 amino acids in the coding sequence constitute the MTS. 2. Make the necessary volume of depurination buffer for each experiment. Denaturing and neutralizing buffers can be prepared in a large volume and stored at room temperature. The pH adjustment of neutralizing buffer should be performed with HCl after dissolving NaCl and Tris in Milli-Q water. Wearing eye protection glasses is recommended when handling HCl and NaOH. 3. DNA fragments with different amounts of bound proteins and circular DNA with different topological turns need to be electrophoresed differentially in EMSA and the DNA-unwinding assay, respectively. Therefore, extra care should be taken for agarose gel preparation. Before microwaving, weigh the flask containing agarose powder and 1× TAE buffer and record the weight. Microwave the solution for 30 s, take out the flask from the microwave and mix the solution by rotating the flask horizontally. Repeat this procedure until the solution is boiling hot. Handle the flask carefully so as not to induce sudden boiling. Wait for a couple of minutes and carefully but adequately mix the solution by horizontal rotation. Repeat the boiling and mixing a couple of times to completely dissolve the agarose. Wearing eye protection glasses is recommended. When mixing, do not introduce excess bubbles in the gel solution, as it results in a gel with bubbles after setting. Such bubbles interfere with electrophoresis. After the final boiling and mixing, let the temperature of the gel solution decrease, with occasional mixing by horizontal rotation. During this step, weigh the flask and calculate the amount of water evaporated during microwaving. Add Milli-Q water to recover the initial weight of the flask with agarose powder and 1× TAE buffer and mix the gel solution carefully but adequately so that it is uniform. This step is important for agarose gel preparation with an accurate gel

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percentage at all the time. When making an EtBr gel, EtBr is added to the gel solution at this stage. Pour the gel solution slowly without introducing bubbles in the gel. 4. We used a bicinchoninic acid (BCA) protein assay kit with BSA for standard curve generation. Accurate protein concentration is important to obtain consistent results in in vitro assays between trials with different preparation batches of TFAM. The same principle applies to the plasmid DNA concentration. 5. It is important that the majority of the plasmid DNA in your preparation is supercoiled and the preparation is not contaminated by bacterial genomic DNA. The supercoiled plasmid must be free from DNA nicks and so suitable for the DNA-unwinding assay. If a substantial proportion of your plasmid DNA is open circular and/or linear, we recommend starting a new plasmid preparation. 6. When we performed EMSA, after incubating 3 kb DNA fragments with TFAM to allow for DNA-binding, the proteinbound DNA fragments were run in agarose gels and visualized with a DNA-intercalating dye. Because protein-bound DNA fragments have a larger molecular weight than fragments without protein binding, the former migrate slower than the latter in a gel, and the amount of proteins bound to the fragments affects the extent of retardation. 7. We used pBluescript KS(-) here, but other plasmids or PCR-amplified DNA fragments can be used as well. 8. We recommend shaking by hand rather than vortexing to avoid inducing DNA nicks in the plasmid. It is essential that the water phase be recovered without organic-phase contamination during both phenol and chloroform-isoamyl alcohol extraction. These chemicals could inhibit TFAM’s DNA-binding activity if carried over to assay reactions. Careful, slow extraction of the water phase by a micropipette is required. In addition, do not try 100% recovery of the water phase, but instead leave ~5% at each step to avoid organic-phase contamination. 9. After addition of 3 M NaOAc and mixing, and before addition of 100% EtOH, a commercially available carrier (Ethachinmate) can be added to ensure efficient ethanol precipitation of the digested plasmid. This is not essential but can be sometimes helpful. 10. It is important to remove 70% EtOH completely from the microtube so that the DNA pellet dries properly. DNA occasionally does not form a pellet at the bottom of the tube, but spreads on the wide area of the inside wall of tubes. In this case, make sure that you dissolve all the pellets.

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11. To construct the reaction buffer, we used 10× DNA topoisomerase I buffer, supplied with calf thymus DNA topoisomerase I, comprising 350 mM Tris-HCl pH 8.0, 720 mM KCl, 50 mM MgCl2, 50 mM DTT, and 50 mM spermidine. In our experience, in the course of optimizing the experimental conditions, addition of 10 mM EDTA blocked DNA nicking activity, which appeared to be residually present in recombinant TFAM preparations. 12. If you prepare several agarose gels at a time and store them at 4 °C until use, ensure that the gels are at room temperature before loading samples and electrophoresis. 13. We believe that the electrophoresis conditions are important to obtain reproducible results in EMSA. The distance between the electrodes is ~13 cm in our electrophoresis tank. With this information, the electrophoresis current can be adjusted for your apparatus. We believe that the length of the gel (6 cm) is appropriate for this assay. Stop the electrophoresis when the supercoiled plasmid (3 kb), which is run as a control, migrates close to the bottom edge of the gel. It runs just below 2000 bp fragments in the DNA ladder (Fig. 1b). In the case of DNA-unwinding assays, stop the electrophoresis when the supercoiled form reaches around three-fourths of the gel (Fig. 2b). You may need to perform pilot experiments to determine when you stop gel electrophoresis with your apparatus. 14. The bottom of the container to rock gels should be smooth and flat to avoid gel cracking during rocking. Additional rocking of the gels in Milli-Q water for 1 min after the staining step might help reduce the background for DNA visualization. 15. First, the supercoiled plasmid is incubated with topoisomerase I to be converted into its relaxed form, which is then incubated with TFAM in the presence of topoisomerase I. TFAM binding introduces negative supercoiling and the DNA is a closed circular plasmid, which is not free from topological constraints, so topoisomerase I detects and releases the constraints introduced by DNA-binding. 16. EtBr induces topological changes in DNA by intercalating the double helix. If relaxed circular DNA with no nick is electrophoresed in an EtBr-containing gel, it is transformed into a supercoiled form (Fig. 2c). However, if plasmid DNA receives a nick at even a single site during the assay (probably from contaminated nucleases in protein preparation), the plasmid DNA will not be converted to the supercoiled form in the EtBr-containing gel. 17. DNA topoisomerase I for the TFAM DNA-unwinding assay must be a eukaryotic topoisomerase I. E. coli topoisomerase I,

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which is also available commercially, cannot be used as it only relaxes negative supercoils, while the eukaryotic counterpart can relax both negative and positive supercoils (see Fig. 1 in [11]). In our experience, optimization of the amount (units) of topoisomerase I and plasmid DNA added to the reaction and the reaction volume appear to be the key to successfully establishing the assay system. If using DNA topoisomerase I from a different company and/or a different plasmid DNA, you may need to optimize their amounts. 18. For DNA fixation onto the membrane, we used the ultraviolet fixation method. Fixed membranes can be kept at room temperature for a couple of weeks, but we recommend that you proceed to Southern hybridization immediately or within a few days. 19. For Southern hybridization, we used AlkPhos Direct Labelling Reagents (GE Healthcare), following the manufacturer’s instructions, and detected the plasmid DNA clearly using an ImageQuant LAS4000 instrument.

Acknowledgments This work was supported in part by Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science [JSPS KAKENHI grant numbers: JP17K07504 to T.Y., 17H01550 to K.D.] References 1. Kang D, Hamasaki N (2005) Mitochondrial transcription factor A in the maintenance of mitochondrial DNA: overview of its multiple roles. Ann N Y Acad Sci 1042:101–108. https://doi.org/10.1196/annals.1338.010 2. Gustafsson CM, Falkenberg M, Larsson NG (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160. https://doi.org/10. 1146/annurev-biochem-060815-014402 3. Hillen HS, Morozov YI, Sarfallah A, Temiakov D, Cramer P (2017) Structural basis of mitochondrial transcription initiation. Cell 171(5):1072–1081 e1010. https://doi. org/10.1016/j.cell.2017.10.036 4. Takamatsu C, Umeda S, Ohsato T, Ohno T, Abe Y, Fukuoh A, Shinagawa H, Hamasaki N, Kang D (2002) Regulation of mitochondrial D-loops by transcription factor A and singlestranded DNA-binding protein. EMBO Rep 3(5):451–456. https://doi.org/10.1093/ embo-reports/kvf099

5. Wang YE, Marinov GK, Wold BJ, Chan DC (2013) Genome-wide analysis reveals coating of the mitochondrial genome by TFAM. PLoS One 8(8):e74513. https://doi.org/10.1371/ journal.pone.0074513 6. Kang D, Kim SH, Hamasaki N (2007) Mitochondrial transcription factor A (TFAM): roles in maintenance of mtDNA and cellular functions. Mitochondrion 7(1–2):39–44. https:// doi.org/10.1016/j.mito.2006.11.017 7. Kukat C, Wurm CA, Spahr H, Falkenberg M, Larsson NG, Jakobs S (2011) Super-resolution microscopy reveals that mammalian mitochondrial nucleoids have a uniform size and frequently contain a single copy of mtDNA. Proc Natl Acad Sci U S A 108(33): 13534–13539. https://doi.org/10.1073/ pnas.1109263108 8. Ngo HB, Lovely GA, Phillips R, Chan DC (2014) Distinct structural features of TFAM drive mitochondrial DNA packaging versus transcriptional activation. Nat Commun 5:

Assessment of TFAM’s DNA-Binding 3 0 7 7 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncomms4077 9. King GA, Hashemi Shabestari M, Taris KH, Pandey AK, Venkatesh S, Thilagavathi J, Singh K, Krishna Koppisetti R, Temiakov D, Roos WH, Suzuki CK, Wuite GJL (2018) Acetylation and phosphorylation of human TFAM regulate TFAM-DNA interactions via contrasting mechanisms. Nucleic Acids Res 46(7):3633–3642. https://doi.org/10.1093/ nar/gky204 10. Ohgaki K, Kanki T, Fukuoh A, Kurisaki H, Aoki Y, Ikeuchi M, Kim SH, Hamasaki N, Kang D (2007) The C-terminal tail of mitochondrial transcription factor a markedly strengthens its general binding to DNA. J Biochem 141(2):201–211. https://doi.org/10. 1093/jb/mvm020 11. Fang Y, Akimoto M, Mayanagi K, Hatano A, Matsumoto M, Matsuda S, Yasukawa T, Kang

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D (2020) Chemical acetylation of mitochondrial transcription factor A occurs on specific lysine residues and affects its ability to change global DNA topology. Mitochondrion 53:99– 108. https://doi.org/10.1016/j.mito.2020. 05.003 12. Farge G, Mehmedovic M, Baclayon M, van den Wildenberg SM, Roos WH, Gustafsson CM, Wuite GJ, Falkenberg M (2014) In vitroreconstituted nucleoids can block mitochondrial DNA replication and transcription. Cell Rep 8(1):66–74. https://doi.org/10.1016/j. celrep.2014.05.046 13. Fukuoh A, Kang D (2009) Methods for assessing binding of mitochondrial transcription factor A (TFAM) to DNA. Methods Mol Biol 554:87–101. https://doi.org/10.1007/9781-59745-521-3_6

Chapter 12 Identification of Proximity Interactors of Mammalian Nucleoid Proteins by BioID Mari J. Aaltonen and Hana Antonicka Abstract Mitochondrial nucleoids are compact nucleoprotein complexes, in which mtDNA is located, replicated, and transcribed. Several proteomic approaches have been previously employed to identify nucleoid proteins; however, a consensus list of nucleoid-associated proteins has not been generated. Here we describe a proximity-biotinylation assay, BioID, which allows identification of proximity interactors of mitochondrial nucleoid proteins. It uses a promiscuous biotin ligase fused to a protein of interest which covalently attaches biotin to lysine residues of its proximal neighbors. Biotinylated proteins can be further enriched by a biotinaffinity purification and identified by mass-spectrometry. BioID can identify transient and weak interactions and can be used to identify changes in the interactions upon different cellular treatments, for different protein isoforms or for pathogenic variants. Key words mtDNA, Nucleoid, Proximity-dependent biotinylation, BioID, Proximity interactors

1 1.1

Introduction Background

Mammalian mtDNA is a circular 16.5 kb-long molecule, which can be visualized in cells as distinct foci, called nucleoids. The principal protein component of the mammalian nucleoid is the mitochondrial transcription factor TFAM, which binds mtDNA with high affinity (reviewed in [1]). Other proteins associating with mtDNA include the components of the mitochondrial DNA replication (POLG, POLG2, TWNK (Twinkle), SSBP1) and transcription (POLRMT, TFB2M, TEFM) machineries. To understand the function of the mitochondrial nucleoid and its role in mitochondrial biology, it is important to define its protein composition. Mitochondrial nucleoids cannot be easily purified using classical biochemical techniques, and previous attempts at the identification of nucleoid components relied on either enrichment of native or cross-linked nucleoids by gradient fractionation [2, 3] or co-immunoprecipitation of the core mtDNA-interacting

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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proteins [3–6]. Although many of the core components of the mitochondrial DNA replication and transcription machineries were identified by several of these proteomic approaches, these methods also identified a number of other mitochondrial and non-mitochondrial proteins, therefore no consensus inventory could be generated [6]. Proximity-dependent biotinylation is an alternative approach for the identification of mitochondrial nucleoid proteins [7, 8]. Proximity-dependent biotinylation methods typically use biotin ligases or peroxidases (reviewed in [9]). These enzymes are fused to a protein of interest (bait) and the construct is inducibly expressed in cells. Upon addition of a substrate, reactive biotin is covalently attached to the proximal proteins (preys) of the bait in vivo. This leads to biotinylation of protein proximity interactors of the bait, as well as of proteins in the “neighborhood” of the bait. The covalent labeling by biotin permits harsh cell lysis, capture of biotinylated proteins on streptavidin-beads, and subsequent identification of captured proteins by mass spectrometry. The peroxidase APEX [7] and the improved version APEX2 [10] were engineered based on a pea ascorbate peroxidase enzyme. APEX/APEX2 uses H2O2 to oxidize biotin-phenol to biotinphenoxyl radicals. These radicals are short lived and react with electron-rich amino acids (predominantly tyrosine) within an estimated radius of 20 nm [7]. Biotin-phenoxyl radicals can freely diffuse within the environment, thus the use of appropriate controls is necessary, coupled with a SILAC- or a TMT-based ratiometric tagging approach, followed by data filtering centered on the measured ratios. BioID uses a promiscuous bacterial biotin protein ligase BirA*, which catalyses the conversion of biotin to biotinyl-5′-AMP. BirA* has been engineered to create a “cloud” of biotinyl-5′-AMP, which can react with the epsilon-amine group of lysine residues on proximate proteins, usually within a radius of 10 nm [8]. BioID has been extensively used to study the composition and organization of membrane-bound as well as membrane-less organelles (reviewed in [9]). The main difference between APEX and BioID is the length of biotinylation reaction needed for sufficient labeling of neighboring proteins: BirA* requires labeling times between 12 and 24 h, while APEX is capable of biotinylation within minutes. This difference prompted the development of biotin ligases with a higher catalytic activity (TurboID and miniTurbo [11]) that yield robust biotinylation signals in ~10 min and are thus suitable for use in dynamic conditions. 1.2 Considerations for a BioID Experiment Design

In this chapter we describe a BioID protocol for identification of nucleoid-associated proteins using the original BirA* biotin ligase. However, the use of alternate biotin ligases, such as BioID2 (which uses biotin ligase from Aquifex aeolicus, that is smaller in size [12]),

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TurboID, or miniTurbo, should be considered if specific applications are planned. The plasmids for construction of the bait fusion proteins are commonly available from Addgene, or can be obtained from researchers upon request [13]. In general, any nucleoid protein can be used as a bait in BioID experiments; however, the proper localization of the bait to mitochondria needs to be established. Prior evidence of a functional fusion protein with a similarly sized tag (e.g., GFP) can be beneficial. As the majority of mitochondrial matrix proteins contain an N-terminal targeting sequence, C-terminal BirA* fusion is required. For this purpose, a Gateway-cloning compatible pDEST-pcDNA5-BirA*-FLAG-C-ter plasmid [13] is used in this protocol. Overexpression of certain proteins can be detrimental to the cells, thus selection of a clonal cell line should be performed, and where possible, the expression of the bait should be compared to the endogenous protein level. The level of biotinylation by each bait can be monitored by western blotting and it is recommended to be tested prior to mass spectrometry analysis. In the BioID protocol, we use Flp-In™ T-REx™ 293 cells, which enable the creation of stable clones with inducible bait expression. However, many different cell lines can be used, including human primary cells. As the transfection efficiency of human primary cells is very low, doxycycline-inducible lentiviral vectors can be utilized [14]. With a transduction efficiency of 80–90%, pools of transduced cells could be used for BioID experiments, instead of individual clones. The level of the expression of the bait can be adjusted by doxycycline titration to avoid overexpression artifacts as mentioned above. Another important consideration in planning BioID experiments is the use of appropriate controls. In general, it is recommended to use cells without a BirA* construct to monitor endogenously biotinylated proteins, such as mitochondrial carboxylases, as well as cells expressing cytosolic BirA*-FLAG-GFP [13] to detect proteins that become promiscuously biotinylated. In addition to direct proximity interactors, proteins in the neighborhood of the bait are also biotinylated, so it is important to assess the biotinylation of the matrix environment, by targeting BirA* to the matrix. Recently, we used three different mitochondrial-targeting sequences (MTS-COX8, MTS-COX4I1, MTS-OTC) to target BirA* to the mitochondrial matrix [15]. These baits mostly identified the same set of preys; however, we detected some differences between them, suggesting an MTS-dependent localization of these baits. In this chapter we will use three controls for illustration: FlpIn™ T-REx™ 293 cells alone, BirA*-FLAG-GFP, and MTSCOX8-BirA*-FLAG expressing cells. The timeline of the BioID protocol is outlined in Fig. 1. The indicated mass spectrometry identification is dependent on the

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Fig. 1 A schematic diagram of the timeline of the BioID protocol

protocols and instrumentation within the proteomic facilities available, and these facilities should be contacted prior to the pull-down of biotinylated proteins to coordinate on-bead trypsin digest and mass spectrometry sample preparation. The subsequent data analysis is dependent on the standard protocols of the labs and proteomic facilities, and several data visualization tools are available, e.g., Prohits-Viz [16] (https://prohits-viz.org) or OmicsVolcano [17]. To identify high-confidence proximity interactors, scores for detected preys are calculated based on the spectral counts identified in the experimental dataset as well as in the negative controls (FlpIn™ T-REx™ 293 cells alone, or BirA*-FLAG-GFP cells) using a proteomic analysis software such as SAINT (Significance Analysis of INTeractome) [18, 19]. To distinguish the bait-specific proximity interactors from proximity interactors in the mitochondrial matrix neighborhood, the score for each prey in the bait dataset is compared to the score identified by the matrix-targeted BirA* (using, e.g., Prohits-Viz).

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Materials All solutions and materials need to be prepared with ultra-pure water and sterilized for the use in the cell culture protocols.

2.1

Bait Cloning

1. Phusion™ High–Fidelity DNA Polymerase master mix. 2. PCR purification kit. 3. PCR primers (see design of primers in Subheading 3.1.1). 4. cDNA template. 5. Plasmid pDONR™ 221 (Thermo Fisher Scientific). 6. Plasmid pDEST-pcDNA5-BirA*-FLAG-C-ter (https:// gingraslab.org/resources/). 7. Gateway® BP Clonase® II enzyme mix. 8. Gateway® LR Clonase® II enzyme mix. 9. BsrGI restriction enzyme. 10. TOP10 competent cells. 11. LB agar. 12. LB media. 13. SOC media. 14. Kanamycin. 15. Ampicillin. 16. Plasmid isolation kit. 17. 1% agarose gels.

2.2 Generation of Cells Stably Expressing the Bait

1. Flp-In™ T-REx™ 293 cells (ThermoFisher Scientific). 2. Cell culture dishes: 10 cm plates, 15 cm plates, 6-well plates, 12-well plates, 24-well plates. 3. Cell culture media: high-glucose DMEM, 10% fetal bovine serum, 500 units/mL penicillin, 500 μg/mL streptomycin (see Note 1). 4. Washing solution: Phosphate buffered saline (PBS): 0.01 M phosphate buffer, 2.7 mM potassium chloride, 0.137 M sodium chloride, pH 7.4. 5. Trypsinization solution: 0.05% trypsin, 0.02% EDTA in PBS. 6. Cell-freezing media: high-glucose DMEM, 20% fetal bovine serum, 10% DMSO. 7. Transfection media: Opti-MEM (ThermoFisher Scientific). 8. Selection media: high-glucose DMEM, 10% fetal bovine serum, 500 units/mL penicillin, 500 μg/mL streptomycin, 200 μg/mL hygromycin.

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9. Lipofectamine 2000 (ThermoFisher Scientific). 10. pDEST-pcDNA5-BaitX-BirA*-FLAG. 11. pOG44 (ThermoFisher Scientific). 12. Hygromycin. 13. 12 mm round coverslips (thickness No.1, Fisherbrand). 14. Cryovials. 2.3 Selection of a Clone for Downstream Analysis

1. 10 mg/mL tetracycline in 50% ethanol. 2. PBS. 3. 4% formaldehyde in PBS. 4. 0.05% (v/v) Triton X-100 in PBS. 5. IF blocking solution: 5% (w/v) BSA in PBS. 6. Anti-FLAG primary antibody (Sigma, F1804). 7. Anti-TOMM40 primary antibody (Proteintech, 18409-1-AP) or another antibody against mitochondrial protein raised in rabbit. 8. Goat anti-mouse IgG (H+L) highly cross-adsorbed secondary antibody, Alexa Fluor 488 conjugated (Molecular Probes, A-11029). 9. Goat anti-rabbit IgG (H+L) highly cross-adsorbed secondary antibody, Alexa Fluor 594 (Molecular Probes, A-11037). 10. DAPI. 11. Mounting media.

2.4 Pull-Down of Biotinylated Proteins

1. 10 mg/mL tetracycline in 50% ethanol. 2. 20 mM Biotin (see Note 2 for preparation protocol). 3. Streptavidin Sepharose (GE17-5113-01, beads that need to be pelleted by centrifugation) or Streptavidin M-280 Dynabeads (11205D Invitrogen, magnetic beads used together with a DynaMag magnet (Invitrogen)). 4. Protease inhibitor cocktail. 5. RIPA buffer: 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1% (v/v) NP-40/IGEPAL, 1 mM EDTA, 1 mM EGTA, 0.1% (w/v) SDS, 0.5% (w/v) sodium deoxycholate, 1× Protease inhibitor. 6. Wash buffer: same as RIPA buffer but without sodium deoxycholate and protease inhibitor. 7. Benzonase. 8. TAP lysis buffer: 50 mM HEPES-KOH pH 8.0, 100 mM KCl, 10% glycerol, 2 mM EDTA, 0.1% (w/v) NP-40. 9. 50 mM ammonium bicarbonate, pH 8.

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10. Trypsin protease, MS grade. 11. 10% or 12% SDS-PAGE polyacrylamide gel (homemade gels or gels commercially available from multiple sources, e.g., Biorad, Novex can be used). 12. Protein measurement reagents (e.g., Bradford, Lowry, BCA). 13. 2 × Laemmli sample buffer. 14. Pre-stained protein ladder. 15. Nitrocellulose or PVDF membrane. 16. Ponceau S protein staining solution. 17. Anti-FLAG primary antibody (Sigma, F1804). 18. Anti-biotin primary antibody (Jackson ImmunoResearch Labs, 200-002-211). 19. Peroxidase-AffiniPure goat anti-mouse IgG (H+L) secondary antibody (Jackson ImmunoResearch Labs). 20. Wash solution: 20 mM Tris-base, 140 mM NaCl, 0.1% (v/v) Tween-20. 21. Blocking buffer: 5% (w/v) non-fat dry milk in Wash solution. 22. ECL reagent. 23. Nutator. 24. Centrifugal evaporator.

3 3.1

3.1.1

Methods Bait Cloning

Primer Design

The bait of interest is cloned into the pDEST-pcDNA5-BirA*FLAG-C-ter vector using a Gateway cloning system by either amplification of the bait gene by PCR from an established clone or from a cDNA. Optionally, a Gateway-compatible clone of a bait gene without a STOP codon (ready for a fusion protein cloning) might be available from plasmid repositories (e.g., DNASU, Addgene). Such a clone can be used directly for LR cloning as in Subheading 3.1.2, step 6. 1. Choose a bait of interest and design primers according to Table 1 (see Note 3). Note that the reverse primer does not include the STOP codon. 2. Order primers from a supplier.

3.1.2 4)

Cloning (See Note

1. Amplify a bait gene according to the manufacturer’s recommendations using Phusion™ High–Fidelity DNA Polymerase, gene-specific primers and either a cDNA template or a plasmid containing the desired bait gene. 2. Gel purify the amplified PCR product.

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Table 1 Design of primers for a Gateway-compatible PCR cloning of C-terminal BirA*-FLAG fusion proteins Name

Terminus attB

Addition Sequence-specific

BaitX-F

GGGG

ACAAGTTTG TACAAAAAAGCAGGCT

CCACC ATG (15–20 nucleotides) (Kozak)

BaitX-R

GGGG

ACCACTTTGTACAAGAAAGC TGGGT

C Reverse complement (20–30 (Frame) nucleotides)

3. Clone the PCR product into pDONR™ 221 using the Gateway® BP cloning system, according to the manufacturer’s recommendations. 4. Transform TOP10 competent cells, and plate on LB agar plates containing kanamycin. Pick at least three clones, expand and isolate the plasmid. 5. Check the insertion of the PCR product into the resulting pDONR-BaitX plasmid by digestion with the BsrGI restriction enzyme. Verify the sequence of the insert by Sanger sequencing using M13+/- primers. 6. Clone BaitX into pDEST-pcDNA5-BirA*-FLAG-C-ter using the Gateway® LR cloning system (using the pDONR-BaitX as an entry plasmid), according to the manufacturer’s recommendations. 7. Transform TOP10 competent cells, and plate on LB agar plates containing ampicillin. Pick at least three clones, then expand and isolate the plasmid. 8. Check the insertion of the BaitX into the resulting pDESTpcDNA5-BaitX-BirA*-FLAG plasmid by digestion with BsrGI restriction enzyme. 9. Verify the resulting plasmid by Sanger sequencing using pDEST-pcDNA5-BirA*-FLAG-C-ter and, if necessary, genespecific primers (see Note 5). 3.2 Generation of Cells Stably Expressing the Bait

Through all the steps, cells are grown and expanded in cell culture media in a standard humidified 5% CO2 incubator at 37 °C using standard cell culture procedures (see Note 6). 1. Day 1: Seed Flp-In™ T-REx™ 293 cells at 250,000 cells per well in a 6-well plate in 2 mL of media. 2. Day 2: Transfection (see Note 7). 2.1 In a microcentrifuge tube, mix 200 ng pDEST-pcDNA5BaitX-BirA*-FLAG and 2 μg of pOG44 in 250 μL of 1 × Opti-MEM.

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2.2 In a second microcentrifuge tube, mix 5 μL of Lipofectamine 2000 reagent with 250 μL of 1 × Opti-MEM. 2.3 Mix the Opti-MEM/Lipofectamine solution and the Opti-MEM/plasmids solution together and incubate for 20 min at room temperature. 2.4 Change the media on Flp-In™ T-REx™ 293 cells in a 6-well plate for 2 mL of cell culture media without antibiotics (see Notes 1 and 8). 2.5 Add the DNA-lipid complex to the cells and incubate for 4 h in a cell culture incubator. 2.6 Change the media to standard cell culture media without antibiotics (2 mL of media). 3. Day 3: Passage transfected cells into 10 cm plates following standard cell culture procedures: aspirate media, wash cells with PBS, add 0.5 mL of trypsinization solution per well, and monitor cell detachment. Once the cells detach, add 2 mL cell culture media per well, and transfer cell solution to a 10 cm plate. Top up the media to 10 mL total per plate. 4. Day 4: Change media on 10 cm plates for 10 mL of selection media. 5. Day 6–14: Change selection media every 2–3 days until clear visible clones are present. 6. Day 15: Pick up to 6 clones per construct. Mark individual clones on the cell culture plate. Remove media from cells and add 5 mL of PBS. Place a small microscope into the cell culture hood and place the 10 cm plate on the stage. Watching through 10 × lens, carefully pick individual clones with P-20 pipette, not disturbing surrounding clones. Disperse the picked cells into a well in a 12-well plate with 1 mL of selection media by vigorously pipetting up and down (see Note 9). 7. Day 18: Trypsinize the cells in the same well of the 12-well plate to disperse them. Once the cells are trypsinized, add 1 mL of selection media and mix by pipetting up and down. Leave the cells in the same well and let them grow until the cells reach confluency. 8. Day 22: Plate cells to select one or two clones for further analysis (see Subheading 3.3). Trypsinize cells in each 12-well plate and resuspend them in 1 mL of media. Prepare two coverslips per clone in a 24-well plate. Seed 100 μL of the resuspended cells on each of the coverslips in 0.5 mL of cell culture media. Transfer the remaining 800 μL of resuspended cells to a 10 cm plate for expansion. (see Notes 10 and 11) 9. Freeze-down the selected clone(s). Grow cells on a 10 cm plate until ~90% confluency. Trypsinize cells and resuspend in 10 mL

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of cell culture media. Transfer the cell solution to a 15 mL conical tube, and spin at room temperature for 3–5 min at 1000 rpm (200 g). Aspirate the media and resuspend cells in 3 mL of cell-freezing media. Aliquot into two freezing vials. Freeze at -80 °C and store in liquid nitrogen. 3.3 Selection of a Clone for Downstream Analysis

Individual clones are screened by immunofluorescence and confocal microscopy to confirm the expression of the bait as well as its mitochondrial localization. The selected clone is further validated by immunoblotting to evaluate the level of expression of the bait construct and the biotinylation level (see Subheading 3.4). 1. The day after seeding, cells on coverslips (step 8 in Subheading 3.2) induce the expression of the bait construct by the addition of 1 μg/mL tetracycline to the media on one coverslip. Leave the other coverslip non-induced. 2. The next day, aspirate media and add 500 μL of 4% formaldehyde in PBS. Incubate for 20 min at 37 °C. 3. Wash coverslips 3 times with 500 μL of PBS. 4. Permeabilize cells in 500 μL of 0.05% Triton X-100 in PBS for 15 min at room temperature. 5. Wash coverslips 3 times with 500 μL of PBS. 6. Block coverslips in IF blocking solution for 10 min at room temperature. 7. Dilute antibodies in IF blocking solution at 1:4000 for antiFLAG and 1:2000 for anti-TOMM40. Add 300 μL of diluted antibodies to each coverslip and incubate for 1 h at room temperature. 8. Wash coverslips 3 times with 500 μL of PBS. 9. Dilute Alexa Fluor conjugated secondary antibodies (1:2000) and DAPI (1:2000) in IF blocking solution. Add 300 μL of diluted antibodies to each coverslip and incubate for 30 min at room temperature. 10. Wash the coverslips 3 times with 500 μL of PBS. 11. Mount coverslips on a glass slide and leave to set for a couple of hours before microscopic analysis. 12. Clone selection: select a clone with predominant mitochondrial localization (see Fig. 2), as well as with a minimal expression of the construct in non-induced cells.

3.4 Pull-Down of Biotinylated Proteins

The selected clone is expanded, and cells are harvested for a biotin pull-down on streptavidin beads (see Subheading 3.4.2), followed by mass spectrometry analysis. A small fraction of cells should be analyzed by SDS-PAGE and western blotting before the pull-down to validate the level of expression of the bait construct and its

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Fig. 2 Immunofluorescence images of mitochondrial targeting of TFAM-BirA*-FLAG and SSBP1-BirA*-FLAG constructs. Control Flp-In™ T-REx™ 293 cells, cells expressing BirA*-FLAG-GFP, or MTS-BirA*-FLAG were used as controls. FLAG staining is shown in green, the mitochondrial marker TOMM20 in magenta, and DAPI in blue. Scale bar, 10 μm

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biotinylation activity (see Subheading 3.4.1). Optionally, an additional analysis of proteins after the biotin pull-down by western blot analysis can be performed and is specified in Subheading 3.4.3. 1. Expand the selected clone (step 12 in Subheading 3.3) and BioID control cells (Flp-In™ T-REx™ 293 cells alone, Flp-In™ T-REx™ 293 over-expressing BirA*-FLAG-GFP and MTS-BirA*-FLAG) to 3 × 15 cm plates in duplicate (see Notes 12 and 13). Adjust cells to similar confluency during expansion. If performing the optional western blot (Subheading 3.4.3), grow an additional 15 cm plate for biotin pull-down and elution with Laemmli SDS-PAGE sample buffer. 2. Grow cells to 70% confluency. Change media to 20 mL per plate and add 2 μL tetracycline (final concentration 1 μg/mL) and 50 μL biotin (final concentration 50 μM) to each plate to induce protein expression and biotinylation. (see Note 14) 3. Harvest cells 24 h later. Remove media, wash cells twice with 5 mL PBS per plate, and harvest using a cell scraper in 5 mL of PBS into a 15 mL conical tube. 4. Pellet cells in the conical tube at 1000 rpm (200 g) for 3 min and aspirate PBS. Wash with 5 mL of PBS. 5. Take a small fraction (2%, 100 μL) of each sample into a separate microcentrifuge tube for SDS-PAGE/western blot analysis. Pellet cells in the microcentrifuge tube at 1000 g for 5 min and aspirate PBS. At this step, the cell pellet can be frozen at -80 °C for later use, or continue with step 1 of Subheading 3.4.1. 6. Pellet the remaining cells in the conical tube at 1000 rpm (200 g) for 3 min and aspirate PBS. 7. Resuspend cells in 1 mL of PBS, transfer to a microcentrifuge tube, then pellet cells at 1000 g for 5 min, and aspirate PBS. 8. Weigh pellets on a scale and write down the pellet weights. Typical yield is around 0.2–0.4 g. At this step, the cell pellets can be frozen at -80 °C for later use. 3.4.1 SDS-PAGE and Western Blot

Before the actual pull-down, a small sample of the harvested cells is analyzed by SDS-PAGE and immunoblotted with anti-FLAG and anti-biotin antibodies to confirm expression and biotinylation in cells. Two gels are needed: one blot will serve for identification of the bait itself (with an anti-FLAG antibody), and the other one to assess the extent of biotinylation (with an anti-biotin antibody). 1. Resuspend cell pellets in 50–100 μL RIPA lysis buffer (volume depends on cell pellet size) and incubate for 20 min on ice. (see Note 15) 2. Centrifuge cells at 20,000 g for 15 min at 4 °C.

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3. Transfer supernatant to a new microcentrifuge tube. 4. Determine the protein concentration in the lysates using Bradford, Lowry, or other preferred method. 5. Prepare duplicate samples for running two SDS-PAGE gels. Aliquot 15 μg of protein per sample and adjust all samples to the same volume (a lower or higher protein amount is also fine as long as all samples have the same amount of protein). Add an equal volume of 2 × Laemmli sample buffer. 6. Boil samples at 95 °C in a heating block for 5 min. Briefly spin the liquid down. 7. Load samples onto 10% SDS-PAGE gels and include a pre-stained protein ladder. Separate proteins by electrophoresis until the Laemmli sample buffer dye front reaches the bottom of the gel. (see Note 16) 8. Transfer proteins from the gel to a nitrocellulose or PVDF membrane using, for example, a Trans-Blot® SD semi-dry transfer cell (Bio-Rad). Any alternative semidry or wet transfer systems can be used. 9. To assess the protein loading, rinse membrane with water, incubate membrane in Ponceau S protein staining solution for 1–2 min, and rinse again with water. 10. Wash the membrane with washing solution for 5 min with agitation on a rocker to remove the Ponceau S protein stain. 11. Block the membrane in blocking buffer for 30 min at room temperature with agitation. 12. Incubate one membrane with an anti-FLAG antibody (dilution 1:2000 in blocking buffer) and the second membrane with an anti-biotin antibody (dilution 1:2000 in blocking buffer) overnight at 4 °C with agitation (see Note 17). 13. Wash the membrane three times in washing solution with agitation for 15 min each. 14. Incubate membrane with horseradish peroxidase (HRP)conjugated secondary antibodies at the manufacturer’s recommended dilution for 1 h at room temperature with agitation. 15. Wash the membrane three times in washing solution with agitation for 15 min each. 16. Develop the membrane using enhanced chemiluminescence (ECL) reagents followed by a chemiluminescence detection of choice. See Fig. 3 for representative images (see Note 18). 3.4.2 Biotin Pull-Down on Streptavidin Beads

The local proteomics facility should be consulted prior to the pulldown to confirm which steps will be performed at the facility (pull down or trypsin digest), and to coordinate sample delivery (prior to the pull down, or at steps 12, 14, or 20). As mentioned above,

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Fig. 3 Analysis of bait expression and prey biotinylation. Flp-In™ T-REx™ 293 control cells, and cells expressing BirA*-FLAG-GFP, MTS-BirA*-FLAG, SSBP1-BirA*-FLAG, or TFAM-BirA*-FLAG were induced for 24 h by incubation with biotin and tetracycline. Cells were lysed with RIPA buffer and 15 μg of protein was analyzed by 10% Tris/Tricine SDS-PAGE followed by western blotting. Bait protein expression is visualized with an anti-FLAG antibody and biotinylated prey proteins are visualized with an anti-biotin antibody. Ponceau S (PoS) staining serves as a loading control

specific steps for analysis of proteins captured on streptavidin beads by western blotting are specified in Subheading 3.4.3. 1. Resuspend cell pellets in RIPA buffer at 1:10 (pellet weight in g: lysis buffer volume in mL). 2. Incubate at 4 °C for 1 h on a rotating platform (nutator). Use the same amount of sample material for each sample in the downstream analysis. 3. Add 1 μL of benzonase (250 U) to each sample. 4. Sonicate lysates at 65% amplitude for 3 × 10 s bursts, with 2 s rest in between on ice. 5. Centrifuge for 30 min at 20,000 g at 4 °C.

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6. During the centrifugation, prepare the beads. Add 30 μL of streptavidin beads per sample to a microcentrifuge tube. Use a wide bore or cut tip to allow easy pipetting of the beads. Wash beads 3 times with 1 mL wash buffer. For streptavidin agarose beads, pellet beads at 400 g for 1 min in-between washes and aspirate the supernatant. For streptavidin M-280 Dynabeads, place the microcentrifuge tube with Dynabeads on the DynaMag magnet to attract the beads on the side of the tube and aspirate the supernatant. Resuspend beads in wash buffer (100 μL per 30 μL bed volume). 7. Collect supernatant into a new microcentrifuge or conical tube depending on the volume. If carrying out the optional western blot (Subheading 3.4.3): determine protein concentration using a preferred method. Adjust lysate concentrations and volumes to similar amounts with RIPA buffer and take input samples corresponding to 1% of protein to be used for binding (e.g., adjust protein amount used for binding to 2 mg and take a 20 μg input sample per gel). Add an appropriate volume of 2 × Laemmli SDS-PAGE sample buffer to the input and store at -20 °C. 8. Add 100 μL of resuspended streptavidin beads in wash buffer (corresponds to 30 μL of original bed volume) to each sample. 9. Incubate at 4 °C on a nutator for 3 h to bind biotinylated proteins on the beads. 10. Pellet beads (by centrifugation for 1 min at 400 g for streptavidin beads, or on a magnet for Dynabeads) and remove supernatant. Transfer beads to a 1.5 mL microcentrifuge tube in 1 mL wash buffer. 11. Wash beads by pipetting up and down (4 × per wash step) in wash buffer four times. Pellet beads in-between washes (by centrifugation at 400 g for 1 min for streptavidin beads, or using a magnet for Dynabeads) and aspirate supernatant (see Note 19). For the optional western blot, proceed to Subheading 3.4.3. 12. Wash two times in 1 mL TAP lysis buffer. 13. Wash the beads three times in 1 mL of 50 mM ammonium bicarbonate, pH 8. Pellet the beads in-between washes (400 g, 1 min; or magnet) and aspirate supernatant. After the last wash, aspirate all residual 50 mM ammonium bicarbonate by pipetting. 14. Resuspend the beads in 200 μL of 50 mM ammonium bicarbonate, pH 8 containing 1 μg of trypsin to digest proteins on beads. 15. Incubate samples at 37 °C overnight with mixing on a rotating disc.

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16. The next day, add an additional 0.5 μg of trypsin (in 10 μL of 50 mM ammonium bicarbonate pH 8) to each sample and incubate for an additional 2 h at 37 °C with mixing on a rotating disk. 17. Pellet the beads by centrifugation at 400 g for 2 min and transfer supernatant to a fresh microcentrifuge tube. 18. Wash the beads two times with 150 μL of 50 mM ammonium bicarbonate, pH 8 (by resuspending and then pelleting beads at 400 g for 2 min in-between) and combine the washes with the original supernatant. Centrifuge the pooled supernatant at 16,100 g for 10 min and transfer most of the supernatant, leaving 30 μL to avoid carrying over the beads, to a new 1.5 mL microfuge tube. 19. Dry samples in a centrifugal evaporator. 3.4.3 Western Blotting of Biotinylated Proteins

1. Wash beads two times in PBS. 2. Centrifuge samples at 400 g for 1 min to pellet beads, and carefully aspirate the remaining supernatant. 3. Elute bound proteins by adding 40 μL of 1 × Laemmli SDSPAGE sample buffer and incubate for 5 min in a 95 °C heating block. 4. Pellet beads (by centrifugation at 400 g for 1 min for streptavidin beads, or using a magnet for Dynabeads) and collect eluate to a new microcentrifuge tube. Repeat elution with an additional 40 μL of 1 × Laemmli buffer and combine the eluates. 5. Run samples on SDS-PAGE gels (as described in Subheading 3.4.1) depending on practices in the lab. Boil samples for 5 min in a 95 °C heating block. Load input and leave an empty well before loading the eluate samples. Load 25–50% (20–40 μL) of the eluates. Incubate with the anti-biotin antibody to confirm enrichment of biotinylated proteins (see Note 18). See Fig. 4 for representative images (see Note 20).

4

Notes 1. Cell culture medium used for transfection does not contain antibiotics. 2. Dissolve 100 mg of biotin into 2.04 mL of 28–30% ammonium hydroxide solution. Neutralize with 18 mL of 1N HCl by keeping the biotin solution on ice and adding ~5 mL of HCl every 2–3 min to avoid heating the solution and precipitation of biotin. If the biotin starts to precipitate, titrate back with a drop of the ammonium hydroxide until the biotin redissolves.

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Fig. 4 Analysis of biotin affinity purification by streptavidin beads. Flp-In™ T-REx™ 293 control cells, and cells expressing BirA*-FLAG-GFP, MTS-BirA*FLAG, SSBP1-BirA*-FLAG, or TFAM-BirA*-FLAG were incubated with biotin and tetracycline, and biotinylated proteins were purified by affinity purification with streptavidin beads. Input (1%, corresponding to 15 μg of lysate) and eluate (25%, corresponding to 20 μL of eluate from 1.5 mg of protein input) samples were analyzed on 10% Tris/Tricine SDS-PAGE gels. Biotinylated prey proteins following affinity purification are visualized with an anti-biotin antibody

Filter sterilize and store at 4 °C. The biotin solution is stable for 6–12 months. 3. Primer design is specific for C-terminal fusion of BirA*-FLAG. It is very important to exclude the STOP codon from the reverse primer design. The length of gene-specific nucleotides should be adjusted to achieve similar Tm. As this protocol can also be applied to other mitochondrial proteins, it might be necessary to create an N-terminal BirA*-FLAG fusion, especially for outer mitochondrial membrane proteins. For N-terminal BirA*-FLAG fusion proteins, follow these primer design recommendations: BaitX-F: GGGGACAAGTTTGTA CAAAAAAGCAGGCTTC -ATG--15-20 sequence-specific nucleotides; BaitX-R: GGGGACCACTTTGTACAA GAAAGCTGGGTTCA--20-30 sequence-specific nucleotides. For N-terminal BirA*-FLAG fusion cloning use pDESTpcDNA5- BirA*-FLAG-N-ter plasmid [13]. 4. PCR amplification, PCR product purification, restriction enzyme digestion, verification of plasmid preparations on agarose gels, competent cell transfection, and plasmid preparations

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are performed according to standard laboratory procedures. The individual steps are mentioned in a generalized manner. 5. pDEST-pcDNA5-BirA*-FLAG sequencing primers: C-ter-F: 5′- gtttagtgaaccgtcagatc-3′; C-ter-R: 5′- ggtctggatgtgcttgttg3′; N-ter-F: 5′- agcaggacggcatcatc -3′; N-ter-R: 5′- cacctactcagacaatgcgatgc -3′. 6. Prior to the experiments, test Flp-In™ T-REx™ 293 cells for mycoplasma contamination using MycoAlert (Lonza). Regular mycoplasma testing is recommended to avoid contamination. 7. Alternative transfection reagents can be used according to the manufacturer’s protocols. 8. Flp-In™ T-REx™ 293 cells can detach easily from the plate, thus gentle handling is required, especially during washes of the cells on plates. 9. Alternative methods for picking clones can be used, including cloning rings or an agarose overlay method, depending on the established techniques in each laboratory. Also, single cell clones can be obtained by counting the cells and plating diluted cells on a 96-well plate, using a dilution factor that yields 40–50 cells per plate. 10. At this step, it is recommended to screen all the clones per construct at the same time. If the clones are not all confluent, >50% confluency is sufficient, however scaling up for immunofluorescense analysis (seeding more cells per coverslip) is recommended. Optionally, coverslips can be treated with poly-D-lysine hydrobromide to improve cell attachment. 11. Cells plated on coverslips are further used for validation of mitochondrial localization and bait expression by immunofluorescence analysis (see Subheading 3.3). Based on this analysis, one or two clones are selected for further downstream analysis (see Subheading 3.4). The rest of the clones are either discarded or can be frozen down. 12. Control selection: Cells contain endogenous proteins which use biotin as a cofactor, and most of these are mitochondrial carboxylases. These proteins are also detected using an antibiotin antibody; thus, the non-induced cells are used as a negative control. It is advantageous to include control cells (without a bait construct) for the comparison of the pattern of biotinylation by the bait relative to the endogenously biotinylated proteins. 13. It is recommended to prepare two biological replicates with three plates each for mass spectrometry analysis, and to perform the biotin pull-down in parallel. 14. If several baits are being prepared, it might be advantageous to collect samples for the downstream analysis and freeze the cell

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pellets as they are ready. Once all samples are collected, samples can be analyzed together. 15. Other solubilization buffers can be used, such as NP-40 lysis buffer, 1.5% N-docecyl-β-D-maltoside in PBS, 1% Triton X-100 in PBS, etc. 16. Depending on the size of the protein of interest used as a bait, 8%, 10% or 12% SDS-PAGE gels might be preferentially used, as the BirA*-FLAG tag adds an additional ~36 kDa to the total size of the bait protein. 17. Primary antibodies can be reused many times over long periods of time, when stored in blocking solution at -20 °C. 18. Since the tagged bait will biotinylate itself, the bait will be the most prominent band in the anti-biotin immunoblot. Higher molecular weight proteins contain more lysine residues which can be biotinylated, and thus will appear more prominently in the biotin immunoblot. 19. Optionally, if required, one stringent wash with 2% SDS buffer (2% SDS, 50 mM Tris–HCl pH 7.5) can be performed prior to the washes in wash buffer. This extra step will dissociate most protein complexes and lead to identification of a reduced number of preys. 20. Additional gels can be run to be immunoblotted with antibodies of interest, e.g., presumed interacting partners of the bait.

Acknowledgments We thank Kathleen Daigneault for technical help with experiments in Figs. 2, 3, and 4; and Dr. Eric A. Shoubridge for critical reading of the manuscript. This work was supported by grants from the United Mitochondrial Disease Foundation and the Canadian Institutes of Health Research. References 1. Farge G, Falkenberg M (2019) Organization of DNA in mammalian mitochondria. Int J Mol Sci 20(11). https://doi.org/10.3390/ ijms20112770 2. Bogenhagen DF, Rousseau D, Burke S (2008) The layered structure of human mitochondrial DNA nucleoids. J Biol Chem 283(6): 3665–3675. https://doi.org/10.1074/jbc. M708444200 3. Rajala N, Hensen F, Wessels HJ, Ives D, Gloerich J, Spelbrink JN (2015) Whole cell formaldehyde cross-linking simplifies purification of mitochondrial nucleoids and associated

proteins involved in mitochondrial gene expression. PLoS One 10(2):e0116726. https://doi.org/10.1371/journal.pone. 0116726 4. He J, Cooper HM, Reyes A, Di Re M, Kazak L, Wood SR, Mao CC, Fearnley IM, Walker JE, Holt IJ (2012) Human C4orf14 interacts with the mitochondrial nucleoid and is involved in the biogenesis of the small mitochondrial ribosomal subunit. Nucleic Acids Res 40:6097. https://doi.org/10.1093/nar/gks257. gks257 [pii]

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5. Wang Y, Bogenhagen DF (2006) Human mitochondrial DNA nucleoids are linked to protein folding machinery and metabolic enzymes at the mitochondrial inner membrane. J Biol Chem 281(35):25791–25802. https://doi.org/10.1074/jbc.M604501200 6. Hensen F, Cansiz S, Gerhold JM, Spelbrink JN (2014) To be or not to be a nucleoid protein: a comparison of mass-spectrometry based approaches in the identification of potential mtDNA-nucleoid associated proteins. Biochimie 100:219–226. https://doi.org/10.1016/ j.biochi.2013.09.017 7. Rhee HW, Zou P, Udeshi ND, Martell JD, Mootha VK, Carr SA, Ting AY (2013) Proteomic mapping of mitochondria in living cells via spatially restricted enzymatic tagging. Science 339(6125):1328–1331. https://doi.org/10. 1126/science.1230593 8. Roux KJ, Kim DI, Raida M, Burke B (2012) A promiscuous biotin ligase fusion protein identifies proximal and interacting proteins in mammalian cells. J Cell Biol 196(6):801–810. https://doi.org/10.1083/jcb.201112098 9. Samavarchi-Tehrani P, Samson R, Gingras AC (2020) Proximity dependent biotinylation: key enzymes and adaptation to proteomics approaches. Mol Cell Proteomics 19(5): 757–773. https://doi.org/10.1074/mcp. R120.001941 10. Lam SS, Martell JD, Kamer KJ, Deerinck TJ, Ellisman MH, Mootha VK, Ting AY (2015) Directed evolution of APEX2 for electron microscopy and proximity labeling. Nat Methods 12(1):51–54. https://doi.org/10.1038/ nmeth.3179 11. Branon TC, Bosch JA, Sanchez AD, Udeshi ND, Svinkina T, Carr SA, Feldman JL, Perrimon N, Ting AY (2018) Efficient proximity labeling in living cells and organisms with TurboID. Nat Biotechnol 36(9):880–887. https://doi.org/10.1038/nbt.4201 12. Kim DI, Jensen SC, Noble KA, Kc B, Roux KH, Motamedchaboki K, Roux KJ (2016) An improved smaller biotin ligase for BioID proximity labeling. Mol Biol Cell 27(8):

1188–1196. https://doi.org/10.1091/mbc. E15-12-0844 13. Couzens AL, Knight JD, Kean MJ, Teo G, Weiss A, Dunham WH, Lin ZY, Bagshaw RD, Sicheri F, Pawson T, Wrana JL, Choi H, Gingras AC (2013) Protein interaction network of the mammalian Hippo pathway reveals mechanisms of kinase-phosphatase interactions. Sci Signal 6(302):rs15. https://doi. org/10.1126/scisignal.2004712 14. Samavarchi-Tehrani P, Abdouni H, Samson R, Gingras AC (2018) A versatile lentiviral delivery toolkit for proximity-dependent biotinylation in diverse cell types. Mol Cell Proteomics 17(11):2256–2269. https://doi.org/10. 1074/mcp.TIR118.000902 15. Antonicka H, Lin ZY, Janer A, Aaltonen MJ, Weraarpachai W, Gingras AC, Shoubridge EA (2020) A high-density human mitochondrial proximity interaction network. Cell Metab 32(3):479–497 e479. https://doi.org/10. 1016/j.cmet.2020.07.017 16. Knight JDR, Choi H, Gupta GD, Pelletier L, Raught B, Nesvizhskii AI, Gingras AC (2017) ProHits-viz: a suite of web tools for visualizing interaction proteomics data. Nat Methods 14(7):645–646. https://doi.org/10.1038/ nmeth.4330 17. Kuznetsova I, Lugmayr A, Rackham O, Filipovska A (2021) OmicsVolcano: software for intuitive visualization and interactive exploration of high-throughput biological data. STAR Protoc 2(1):100279. https://doi.org/10. 1016/j.xpro.2020.100279 18. Teo G, Liu G, Zhang J, Nesvizhskii AI, Gingras AC, Choi H (2014) SAINTexpress: improvements and additional features in Significance Analysis of INTeractome software. J Proteome 100:37–43. https://doi.org/10.1016/j.jprot. 2013.10.023 19. Choi H, Larsen B, Lin ZY, Breitkreutz A, Mellacheruvu D, Fermin D, Qin ZS, Tyers M, Gingras AC, Nesvizhskii AI (2011) SAINT: probabilistic scoring of affinity purificationmass spectrometry data. Nat Methods 8(1): 70–73. https://doi.org/10.1038/nmeth. 1541

Chapter 13 Localization of Mitochondrial Nucleoids by Transmission Electron Microscopy Using the Transgenic Expression of the Mitochondrial Helicase Twinkle and APEX2 David Pla-Martı´n, Felix Babatz, and Astrid C. Schauss Abstract Reminiscent of their evolutionary origin, mitochondria contain their own genome (mtDNA) compacted into the mitochondrial chromosome or nucleoid (mt-nucleoid). Many mitochondrial disorders are characterized by disruption of mt-nucleoids, either by direct mutation of genes involved in mtDNA organization or by interfering with other vital proteins for mitochondrial function. Thus, changes in mt-nucleoid morphology, distribution, and structure are a common feature in many human diseases and can be exploited as an indicator of cellular fitness. Electron microscopy provides the highest possible resolution that can be achieved, delivering spatial and structural information about all cellular structures. Recently, the ascorbate peroxidase APEX2 has been used to increase transmission electron microscopy (TEM) contrast by inducing diaminobenzidine (DAB) precipitation. DAB has the ability to accumulate osmium during classical EM sample preparation and, due to its high electron density, provides strong contrast for TEM. Among the nucleoid proteins, the mitochondrial helicase Twinkle fused with APEX2 has been successfully used to target mt-nucleoids, providing a tool to visualize these subcellular structures with high contrast and with the resolution of an electron microscope. In the presence of H2O2, APEX2 catalyzes the polymerization of DAB, generating a brown precipitate that can be visualized in specific regions of the mitochondrial matrix. Here, we provide a detailed protocol to generate murine cell lines expressing a transgenic variant of Twinkle, suitable to target and visualize mt-nucleoids. We also describe all the necessary steps to validate the cell lines prior to electron microscopy imaging and offer examples of anticipated results. Key words Mitochondria, Nucleoid, TEM, APEX2

1

Introduction The mitochondrial genome (mtDNA) is a dynamic doublestranded molecule of 16.5 kb that encodes for 13 proteins, 22 tRNAs, and 2 rRNAs. mtDNA is compacted into a nucleoprotein complex within the mitochondrial matrix, known as the mitochondrial nucleoid (mt-nucleoid) [1]. Each nucleoid contains approximately one copy of mtDNA and is structured in elongated particles of nearly 100 nm [2]. Proteins associated with the

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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mt-nucleoid are involved in a variety of roles such as mtDNA replication, maintenance, and transcription. The major component of mtDNA is the mitochondrial transcription factor A (TFAM), which compacts and protects the DNA molecule. However, other proteins can be temporally associated with the mt-nucleoid, and among them, the mitochondrial helicase, Twinkle, is required for mtDNA strand separation at the replication fork and is essential to maintain mtDNA copy number at physiologic levels [3, 4]. Visualization of mtDNA has historically been linked to the development of microscopy techniques. The discovery of the mt-nucleoid in the mitochondrial matrix took place in 1963 using electron microscopy [5]. The study of the mitochondrial ultrastructure of a chicken embryo using a wide variety of fixatives revealed the presence of intramitochondrial fibers with electron-dense qualities resembling the bacterial nucleoplasm. The advancement of fluorescence confocal microscopy, immunostaining techniques, and chemicals reacting specifically against DNA has improved the ability to detect and visualize mtDNA. In classical confocal microscopy, mt-nucleoids can be observed as particles of different sizes depending on their distribution within the mitochondrial network. However, the limited spatial resolution of light microscopes (200 nm in the x–y axis and > 600 nm in the z) restricts the determination of structures of smaller size. Lately, with the development of super resolution microscopy, this resolution limit has been overcome: for single-molecule localization-based microscopy (SMLM), the resolution can reach 20–40 nm [6] and, with STED (STimulated Emission Depletion) microscopy, the resolution reaches less than 40–50 nm in x–y and 300 nm in z, allowing one to resolve even single mt-nucleoids [2]. The highest resolution of cellular structures is provided by transmission electron microscopy (TEM), reaching an atomic resolution for proteins in cryo-EM, and making it suitable to resolve any cellular structure on a molecular level [7]. The major advantage of electron microscopy, despite the gain in resolution, is the context, showing not only the protein or structure of interest but the surrounding area. Classically, detection of a specific protein with electron microscopy can be achieved using specific antibodies against a protein and secondary antibodies conjugated with electron-dense gold nanoparticles [8]. This labeling strategy provides spatial information and subcellular localization by showing a small black dot near the labeled protein. During sample preparation, however, biological samples are treated with strong fixatives and electron-dense chemicals to deliver contrast. Furthermore, samples must be embedded in resins suitable for ultrathin cutting. These treatments could, ultimately, modify antigen reactivity and impede antibody detection. Immunogold labeling can be also applied onto classical Epon sections, but in this case, most of the epitopes for antibody recognition are covered up and have to be

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retrieved in additional treatments, at the expense of lower ultrastructural preservation. Following this technique, mtDNA visualization by TEM has been achieved, but the resolution of mitochondrial membranes was strongly compromised [9]. In alternative approaches, mild fixatives to maintain antigenicity or less hydrophobic lowicryl embedding (e.g., HM20, LRWhite) can be used to improve antibody recognition [10]. In other immunolabeling procedures such as the Tokuyasu technique, the samples are cut under frozen conditions, leaving out the need for resin embedding and, with that, circumventing the follow-up problem in antigen recognition [11]. However, special equipment for cryosectioning as well as very gentle handling of the fragile frozen sections is needed. Independently of all the discussed embedding strategies, a prerequisite for the detection of any protein by immunogold labeling is the availability of good antibodies with strong reactivity and high specificity. Horseradish peroxidase (HRP)-coupled antibodies have been used for decades in immunohistology. In the presence of H2O2, HRP catalyzes the precipitation of a brown polymer from diaminobenzidine (DAB). This polymer has the ability to accumulate osmium during classical TEM preparation, and due to its high electron density, generates a precipitate visible by TEM [12, 13]. Recently, several genetic tools following this approach have been engineered with the advantage that they can be incorporated into a standard routine in any EM lab. Thus, HRP has been fused to membrane proteins for correlative fluorescence and electron microscopy (CLEM) [14]. HRP provides excellent contrast but requires the presence of Ca2+ ions, restricting its use to some cellular compartments and making it incompatible with others such as cytosol. Moreover, HRP demands special posttranslational modifications such as disulfide bonds and glycosylation, which could interfere with the function of the fusion protein, its transport, and the correct subcellular localization. Alternatively, miniSOG (mini singlet oxygen generator), a fluorescent flavoprotein engineered from Arabidopsis phototropin 2, catalyzes DAB polymerization upon strong illumination with blue light [15]. This reaction, however, demands intense sample lighting and oxygen, requiring special equipment and hindering protocol adaptation. Finally, APEX protein and its direct derivative APEX2 [16, 17], provide stronger contrast due to increased DAB precipitation during EM preparations compared to the other alternatives. APEX2 is a 28 kDa monomeric peroxidase reporter derived from dimeric pea or soybean ascorbate peroxidases. Because of its unique properties, APEX2 can be used for proteomic mapping and for electron microscopy detection at a suborganellar level. After cellular fixation, APEX2 is incubated in the presence of DAB together with H2O2, inducing DAB deposition restricted to the cellular compartment of the fusion protein (Fig. 1a, b). This technique does not

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Fig. 1 Twinkle and APEX2 as a method to visualize mt-nucleoids. (a) Schematic representation of mt-nucleoids and DAB precipitation induced by APEX2 in presence of H2O2. (b) Electron microscope image of wild-type C2C12 and cells expressing mitochondrial-targeted APEX2-V5 and Twinkle-APEX2-V5. Mitochondrial matrix-targeted APEX2 generates a dark signal located specifically in the matrix. In contrast, TwinkleAPEX2-V5 induces the restricted deposition of DAB, revealing the relative position of mt-nucleoids. Arrows indicate the location of mt-nucleoids. DAB precipitate and osmium post-staining were generated following the protocol described in this chapter. Scale bar, 0.5 μm. (c) Time schedule to generate murine cell lines expressing mt-nucleoids APEX2 tagged suitable for electron microscope imaging. Generated with biorender.com

require special equipment, and DAB precipitate can be generated with standard chemical reagents, facilitating its adaptation in any standard cell biology lab [17]. Thus, APEX2 has emerged as a potent alternative to visualize technically any protein of interest into the resolution of an EM. As with any technique involving genetic manipulation of a protein, the use of APEX2 has some limitations to consider. First, validation of protein functionality and cellular localization might need to be carried out, to exclude that protein fusion to the 28 kDa APEX2 interferes with expected protein localization and cellular function. Specific to visualizing mt-nucleoids, Twinkle has been shown to maintain its function when APEX2 is fused to the C-terminus side of the protein [18]. Other nucleoid proteins such as TFAM might also provide the requested specificity, and indeed, TFAM-APEX2 has been expressed in a Drosophila knock-in to analyze mtDNA content and structure [19]. Nonetheless, in human cell lines, TFAM-APEX2 fusion, despite localizing in the mitochondrial matrix, did not co-localize with dsDNA detected within mitochondria, and hence, the proper TFAM function could not be assured, making it inadequate for further analysis [18]. In this chapter, we provide a detailed protocol to generate murine cell lines suitable to visualize mt-nucleoids using Twinkle

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as bait. Additionally, we give a full description of the controls required to conduct a successful experiment and all preliminary steps before imaging in the electron microscope. Finally, we describe the steps to process and stain cellular samples and provide contrast to distinguish the specific staining within the electron microscope.

2

Materials To achieve high levels of expression and high numbers of transfected cells, we recommend transducing the cells with retroviral vectors following the selection of independent clones. All plasmids to generate ecotropic or amphotropic retroviruses can be purchased at the public plasmid repository Addgene. Retroviruses must be generated by co-transfecting the required plasmids in a packaging cell line. A timeline for the required experiments can be found in Fig. 1c.

2.1

Plasmids

1. Retroviral plasmid encoding Twinkle-APEX2-V5 fusion protein: for instance, pBABE-Puro Twinkle-APEX2-V5 (see Note 1). 2. Retroviral helper plasmid: pCL-ECO (see Note 2).

for

ecotropic

retroviruses,

3. Control plasmids: we recommend the use of control plasmids to visualize the different forthcoming steps. Retroviral plasmids encoding a fluorescent protein for transfection/transduction and mitochondrial matrix APEX2 for DAB deposition control can be purchased at Addgene (see Note 3). 4. High purity endotoxin-free plasmid isolation kit. 2.2 Generation of Murine Cell Lines Expressing NucleoidTargeted Transgene (See Note 4)

1. Retroviral packaging cell line: HEK293. Other modified packaging cell lines to avoid the use of helper plasmids can also be found commercially. 2. Murine cell line: many already established cell lines can be purchased. Alternatively, the protocol can also be applied to mouse primary cells. 3. Polyethylenimine (PEI) linear: 1 mg mL-1 solution in ultrapure water (Milli-Q water). Allow complete dissolution by adding HCl to acidify the water. Once the solution is clear, adjust the pH to 7.0 with NaOH. Filter through a 0.22 μm membrane, aliquot, and store at -80 °C. Working aliquots can be stored at 4 °C for up to 2 months. 4. Opti-MEM: reduced serum media with non-phenol red.

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5. Supplemented DMEM: 4.5 g L-1 glucose, 10% inactivated FBS, 1× penicillin/streptomycin, 1× glutamine or another appropriate cell-growing medium. 6. Polybrene (hexadimethrine bromide) 8 mg mL-1: dissolve 80 mg in water. Filter through a 0.22 μm membrane and store at 4 °C. 7. PES or nylon syringe filters 0.45 μm. 8. T25 and T75 flasks. 2.3 Validation of Cell Clones by Immunofluorescence

1. Specific antibody to detect Twinkle tagged version (see Note 5). 2. Anti-mitochondrial membrane protein or anti-mitochondrial matrix: different antibodies can be found commercially. We obtain optimal results with rabbit anti-TOM20 and rabbit anti-LRPPRC from Proteintech. 3. dsDNA monoclonal antibody: we recommend mouse monoclonal anti-dsDNA purchased from Abcam (Clone 35I9 DNA). 4. Biotin-phenol 500 mM: dissolve 91 mg in 0.5 mL DMSO. Aliquot and store at -80 °C. 5. H2O2 1 M: provided as a 30% solution, dilute 510 μL of 30% H2O2 in 5 mL water. 6. Trolox 100 mM: dissolve 1 g in 40 mL DMSO. Store at -20 ° C. 7. Sodium ascorbate 1 M: prepare fresh in ddH2O. 8. Sodium azide 1 M: prepare fresh in ddH2O. 9. Phosphate buffered saline (PBS). 10. 0.1% Triton X-100. 11. 10% Normal goat serum in PBS. 12. DAPI-Fluoromount mounting medium. 13. Quencher solution: 10 mM Na-ascorbate, 10 mM NaN3, 5 mM Trolox in PBS. 14. 4% PFA/PBS: dissolve 20 g of paraformaldehyde in 400 mL PBS. Increase the pH to 11 with NaOH and let it dissolve. Once the solution is clear, adjust to pH 7.4 with HCl. Adjust the volume to 500 mL with PBS. Aliquot and freeze at -20 °C. When stated, this may contain 10 mM Na-ascorbate, 10 mM NaN3, and 5 mM Trolox. 15. Fluorescent NeutrAvidin protein. 16. Anti-rabbit and anti-mouse fluorescent secondary antibodies.

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1. Aclar foil disc: cut with a hole puncher and sterilize using UV light. 2. 2% Glutaraldehyde, 2% sucrose in HEPES pH 7.4: fixative should be prepared freshly the same day. 3. 0.1 M Cacodylate buffer: adjust pH to 7.3. 4. 0.1 M Glycine: prepare in 0.1 M cacodylate buffer. 5. Diaminobenzidine: prepare 0.5 mg mL-1 in 0.1 M cacodylate buffer. Dissolve 10 mg diaminobenzidine tablets in 20 mL of 0.1 M cacodylate buffer. Filter through a sterile filter to remove undissolved DAB. Add 20 μL of 30% H2O2 to achieve a final concentration of 0.03% right before use. 6. 1% Osmiumtetroxid: diluted from 4% aqueous stock solution. 7. 1.5% Potassium hexacyanoferrate(III): dissolved in ddH2O. 8. Ethanol series for dehydration: prepare 50%, 70%, 90%, and 100% in ddH2O. Store at -20 °C. 9. Epon: For 20 g Epon, mix 10 g epoxy embedding medium with 5.5 g epoxy embedding medium hardener DDSA and 4.5 g epoxy embedding medium hardener MNA. Add 0.4 g accelerator DMP30. 10. 8 mm polyethylene flat embedding capsules (TAAB). 11. Ultramicrotome provided with Diamond Knife 35° (Diatome): align the block face parallel to the knife edge to collect as many sections from the cell monolayer as possible. 12. 1.5% uranyl acetate: dissolve uranyl salt in ddH2O and store at 4 °C. 13. Lead citrate solution: mix 800 μL ddH2O with 155 μL 1.3 M tri-sodium citrate and 100 μL lead nitrate. To dissolve precipitate, add 200 μL 1 M NaOH. Centrifuge for 10 min at full speed before use. 14. Transmission electron microscope.

3

Methods

3.1 Generation of Ecotropic Retroviruses (See Note 6)

1. Before starting with cell culture protocols, isolate all the required plasmids using Midi scale high purity endotoxinfree kits. 2. Grow HEK293 cells or another packaging cell line in supplemented DMEM in a T75 flask until reaching confluency (see Note 6). Typically, a confluent T75 flask contains approximately 106 cells. Wash with PBS, then trypsinize the cells and count with a haematocytometer. Seed 2.5 × 106 cells in a petri dish p100. Supplement with new medium and allow the cells to attach until next day.

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3. The following day, thaw a new batch of the target cells. For transfection of HEK293 cells, combine 1 mL Opti-MEM, 5 μg pCL-ECO, 10 μg of retrovirus transfer vector, and 40 μL PEI in an Eppendorf tube. Mix and incubate for 15 min at room temperature. Use this time to replace the HEK293 cells medium with 10 mL of new complete DMEM (see Note 7). 4. Add the plasmid–PEI mixture to the HEK293 cells drop by drop, and return the cells to the incubator for 24 h. 5. After this time, replace the HEK293 medium with 10 mL of fresh complete DMEM, and incubate the cells for a further 24 h. 6. Trypsinize the target cells and seed an appropriate number of cells in a T25 flask, to achieve a confluence of 40–50%. 3.2 Generation of Cell Lines Stably Expressing Transgenes

1. Two days after transfection, using the cells transfected with the plasmid encoding a fluorescent protein, check whether transfection was achieved using a fluorescence microscope. 2. Recover the supernatant medium of HEK293 and transfer to a clean 15 mL tube. Filter the supernatant through a 0.45 μm sterile filter and transfer to a new tube. Add polybrene at a final concentration of 8 μg mL-1. Replace the medium of the target cells with the medium containing viruses. Return the cells to the incubator and let them rest for 24 h (see Note 8). 3. The day after transduction, remove the medium containing the viruses and add new complete DMEM. Let the cells grow for at least 48 h (see Note 9). 4. Using a fluorescence microscope, verify that the cells transduced with the fluorescent control plasmid, show the correspondent fluorescence. 5. Start the selection with complete DMEM containing 2.5 μg mL-1 puromycin (see Note 10). Expand the cells and freeze as soon as enough material has been achieved. 6. After 48–72 h of selection, start isolating independent clones using clonal filters or a serial dilution method (see Note 11). 7. After 1 week, scan plates to find single colonies. Discard all the clones derived from more than one cell. Expand the selected clones, freeze, and store in liquid nitrogen.

3.3 Validation of the Cell Lines 3.3.1 mt-Nucleoid Localization of TwinkleAPEX2-V5

1. Seed the cells onto glass coverslips and let them attach. 2. Fix the cells in 4% PFA/PBS for 15 min at room temperature. Wash with PBS and store at 4 °C until use. 3. Transfer the coverslips to a humid chamber. Permeabilize the cells with PBS containing 0.1% Triton X-100 for 30 min. Block with 10% normal goat serum in PBS for 1 h at room temperature.

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Fig. 2 mt-nucleoid localization of Twinkle-APEX2-V5 in murine cells. Prior to analysis of mt-nucleoids by TEM, it is advisable to verify that transgenic expression of Twinkle does not interfere with the expected cellular localization of the protein. α-V5 is used to detect transgenic Twinkle, α-dsDNA to detect mt-DNA, and α-TOM20 as a mitochondrial marker. (a) C2C12 mouse myoblast. (b) MEFs (mouse embryonic fibroblast). Scale bar, 10 μm. Images were acquired using either an (a) Ultraview Vox Spinning Disk microscope or (b) Leica SP5 confocal microscope

4. Incubate the primary antibodies diluted in blocking buffer in a humid chamber (see Note 12). 5. Wash three times with PBS for 10 min each time. Incubate the fluorescent secondary antibodies for 1 h at room temperature in blocking buffer. Wash three times with PBS for 10 min each and mount with DAPI-Fluoromount. 6. Scan in a confocal fluorescence microscope for the desired fluorescence (see Note 13). See Fig. 2 for anticipated results.

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3.3.2 Proximity Biotinylation

APEX2 functionality can be easily assayed by performing a biotin ligation protocol followed by conventional immunofluorescence. 1. Seed the cells onto glass coverslips and let them attach. 2. Incubate with 2.5 mM biotin-phenol for 8 h in complete DMEM (see Note 14). 3. Add H2O2 to a final concentration of 1 mM. Incubate for 1 min at 37 °C. 4. Wash 3× with Quencher solution. 5. Fix with 4% PFA/PBS containing 10 mM Na-ascorbate, 10 mM NaN3, and 5 mM Trolox. 6. Wash 3× with PBS. 7. Proceed with conventional immunofluorescence for the detection of mitochondrial proteins and transgenic Twinkle. To detect biotinylated proteins, incubate NeutrAvidin together with the fluorescent secondary antibodies for 2 h at room temperature (see Note 15). See Fig. 3a for anticipated results.

Fig. 3 Validation of APEX2 activity by light microscopy. Murine myoblast C2C12 cells were transduced with Twinkle-APEX2-V5 plasmid. (a) Confocal image of proximity biotinylation experiment performed as described in this chapter. NeutrAvidin-488 was used to detect biotinylated proteins, α-V5 to detect transgenic Twinkle, and α-TOM20 as a mitochondrial marker. Scale bar, 10 μm. (b) Bright field image of wild-type and transduced C2C12 cells. DAB deposition was induced as described in this chapter. Twinkle-APEX2-V5 transduced cells show a brown signal resembling the mitochondrial pattern. Scale bar, 100 μm

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1. Grow the cells on small discs of aclar foil previously sterilized by UV. 2. Fix for 1 h in 2% glutaraldehyde with 2% sucrose in 0.1 M HEPES buffer pH 7.4. 3. Wash two times with 0.1 M cacodylate buffer for 5 min. 4. Quench the free aldehyde groups by incubating with 0.1 M glycine in 0.1 M cacodylate buffer twice for 20 min each time. 5. Incubate for 30 min in 0.1 M cacodylate buffer containing 0.5 mg mL-1 diaminobenzidine and 0.03% H2O2. 6. Evaluate cells in a bright field microscope for brown precipitate (Fig. 3b).

3.5 Ultrathin Sample Preparation for EM

1. Wash cells briefly three times with 0.1 M cacodylate buffer. 2. Incubate with 1% osmiumtetroxid and 1.5% potassium hexacyanoferrate for 30 min at 4 °C. 3. Wash three times for 5 min with ddH2O and dehydrate the samples using an ascending ethanol series (50%, 70%, 90%, 100%) for 5 min each time at 4 °C. 4. Infiltrate with a mixture of 50% Epon/ethanol overnight at 4 ° C. 5. Infiltrate with pure Epon for two times for 2 h. 6. Place the aclar disc with cells facing upward onto the bottom of a flat TAAB capsule, fill it with Epon, and cure for 48 h at 60 ° C. 7. Remove the TAAB capsule and aclar foil using a razorblade. 8. Cut 70 nm ultrathin sections with an ultramicrotome equipped with a diamond knife. Pick up the sections onto 75 mesh grids or single slot grids and let them dry. 9. Post stain the sections with 1.5% uranyl acetate for 20 min at 37 °C and 3 min with lead citrate solution. 10. Acquire images in a TEM (Fig. 4).

Fig. 4 TEM images of C2C12 wild type and transduced with Twinkle-APEX2-V5. DAB deposition and sample treatment were performed as described in this chapter. Arrows signal accumulation of osmium induced by DAB deposition, and hence, mt-nucleoids. Scale bar, 0.5 μm

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Notes 1. Twinkle has to be engineered to fuse APEX2 in the C-terminus side of the protein. Additionally, to detect the transgene and verify that its expression is in the expected cellular compartment, it is advisable to include a sequence encoding a protein tag. To achieve high levels of cells expressing the transgene and facilitate further experiments, it is advisable to transduce the cells using retroviruses followed by antibiotic selection. Lentiviral plasmids containing the human sequence of Twinkle fused with APEX2-V5 are available at Addgene. For the examples shown in this chapter, we have developed a Mus musculus variant of Twinkle fused with APEX2-V5 in the C-terminus region. 2. Transduction of murine cell lines requires the generation of ecotropic retroviruses. Typically, a transfer plasmid based on MoLV or MSCV murine viruses and a helper plasmid are required. Here, we use the transfer plasmid pBABE-Puro [20] and the helper pCL-ECO [21]. pCL-ECO plasmid contains the viral proteins gag, pol, and env together in one insert and produces helper free and high titer of retroviruses. Other plasmids such pUMVC can be used, but in this case, the ecotropic envelope protein must be provided in another plasmid. Retroviruses are useful when the target cells are dividing cells. Targeting slow-dividing cells or non-murine cell lines will require the use of other helper plasmids to generate amphotropic viruses, also known as lentiviruses. In that case, transfer plasmids can be based on HIV-1 virus and a higher level of security is required. The tropism of the env protein derived from the vesicular stomatitis virus (VSV-G) provides a wide spectrum of potentially infectable cells, including human cells. In that case, the gag, pol, and env proteins are delivered in different plasmids. Normally, it is not possible for a virus generated with this system to produce more viruses after the initial infection. In addition, many lentiviral transfer vectors are selfinactivating vectors containing deletions and insertions in the LTR regions, which interfere with the transcription of complete viruses. 3. The protocol described here contains several critical steps for which we advise the use of control plasmids to visualize the different processes. As a positive control for transfection/transduction, we recommend using a retroviral plasmid encoding a fluorescent protein and verify, under a fluorescence microscope, that transfection/transduction has occurred; for instance, pBABE-Puro vector encoding GFP can be provided

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by different commercial suppliers and public repositories. Any other retroviral plasmid encoding a fluorescent reporter compatible with the helper virus can also be used as a control; for example, we use a retroviral vector pMX2 encoding mCherry fluorescent protein [22]. To visualize APEX2 enzymatic activity in light or electron microscopy, a plasmid encoding APEX2V5 targeted to the mitochondrial matrix or other mitochondrial compartments can be purchased at Addgene. 4. Here, we describe the generation of clones in the murine myoblast cell line C2C12 and mouse embryonic fibroblasts (MEFs). Any other murine cell line or primary culture from a mouse model can be used following the same protocol, but it might require some adaptation. Targeting mt-nucleoids for other species might require some protocol adjustments. We cannot guarantee that murine or human Twinkle is suitable for cells derived from other organisms besides Mus musculus or Homo sapiens, respectively. 5. We use V5 as a tag to detect transgenic Twinkle. Plasmids expressing this fusion protein can be found commercially (see Note 1). Additionally, mouse monoclonal and rabbit polyclonal antibodies from different sources can also be purchased from several suppliers. 6. Generation of ecotropic or amphotropic viruses requires the use of S2 biosecurity level. Please follow the instructions of your employer and local regulation about how to work and discard the items in contact with S2 biohazard material. 7. HEK293 cells are extremely sensible to movements and detach easily. Handle with care to avoid detachment. Avoid rinsing with PBS or adding medium directly on the cells, as this will cause cell detachment and poor results. In the case of using commercially available packaging cell lines already transformed to express endogenously viral proteins, avoid the transfection of pCL-ECO or other helper plasmids. 8. Some protocols related to retroviral infection describe a double step protocol infecting for two consecutive days. We found that only one infection round is enough to achieve proper expression levels of Twinkle without producing overexpression artifacts. However, if low levels of transgene expression are observed, a second round of infection is also possible. For that, after collecting the first viral supernatant, supplement transfected HEK293 with fresh complete DMEM medium and incubate for 24 h more. The next day (72 h after transfection), process supernatant medium as described and reinfect target cells. Alternatively, supernatant can be frozen at -80 °C after filtering to use in future experiments. When required, thaw the viruses-containing medium on ice. The medium will

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carry a lower level of retroviruses, but despite this, it will be still possible to infect cells, although with lower efficiency than freshly collected retroviruses. All material in contact with retroviruses such plates and pipettes should be discarded following local regulations. 9. To allow the genomic insertion of the transgene, cells need to grow for at least 2 days. We found that starting the selection before this time induces cell death and complicates the selection of positive clones. During this time, it is necessary to transfer the cells to a bigger growing surface and expand them. 10. The concentration of puromycin for selection should be determined empirically for each cell line. We found that for C2C12, 2.5 μg mL-1 puromycin is enough to kill all non-transduced cells in 48 h. Other cell lines like MEFs are more resistant to puromycin and required 5 μg mL-1 of antibiotic for selection. These values can be used as a guide, and a dose-response experiment should be performed preliminary to the selection procedure. 11. Transduction with retroviruses might produce different levels of transgene overexpressing cells. We recommend isolating monoclonal lines to achieve consistent results. Some cell lines do not grow well when seeded at low density, so increasing the serum to 20% might be beneficial to generate monoclonal cell lines. This step can be skipped, but the researcher must be aware that some cells might show artifacts and mitochondrial abnormalities due to overexpression of Twinkle. 12. Incubation times and concentration of the antibodies should be determined empirically. The use of different anti-tag antibodies is required, in combination with other mitochondrial markers, to verify that Twinkle is properly expressed. Here, we use V5 rabbit polyclonal and mouse monoclonal antibodies to detect the transgenic Twinkle-APEX2-V5, combined with the mouse monoclonal dsDNA and rabbit polyclonal mitochondrial markers. We found that the best results are from incubating primary antibodies overnight at 4 °C. Clone 35I9 mouse monoclonal dsDNA gives the best result with 1:1000 dilution. Polyclonal antibodies against mitochondrial markers can be used at a 1:500 dilution. Fluorescent secondary antibodies might be incubated at a 1:1000 dilution. 13. Typically, dsDNA should stain both nucleus and mtDNA. mtDNA staining is found as a puncta pattern inside mitochondria (Fig. 2). The strength of nuclear staining is dependent on the cell line or antibody batch, but the suggested monoclonal antibody provides nuclear staining. Twinkle-APEX2-V5 should present a puncta pattern correlative to dsDNA antibody and co-localized with mitochondrial markers. Exacerbated

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expression of the transgene produces continuous V5 staining through the mitochondria. A lower level of expression clones might be selected. If required, dilution of the medium containing retroviruses used for transduction can be used to achieve lower levels of overexpression. 14. We found this concentration and incubation time for C2C12 the best to achieve a high number of biotinylated cells (Fig. 3a). Biotin-phenol concentration and incubation time for other cell lines should be determined empirically. In addition, APEX2 activity requires the presence of heme b cofactor, which is produced endogenously in most organisms. However, in some cell lines, it might be necessary to supplement it. Concentration and incubation time should be determined empirically for such cases. Controls lacking either H2O2 or biotin should be performed as well. 15. NeutrAvidin fluorescent protein can be incubated at 1:500. Detection of biotinylated proteins is necessary to confirm that the activity of APEX2 is restricted to the mitochondria. Due to the resolution of confocal microscopy and the strength of the enzymatic activity, the fluorescence signal derived from NeutrAvidin might not be restricted to nucleoids, but it must colocalize with dsDNA or mitochondrial markers.

Acknowledgments This work was supported by grants from the Deutsche Forschungsgemeinschaft (PL 895/1-1) and Ko¨ln Fortune (341/2019) to DPM and CRC1218 to FB and ACS. References 1. Stewart JB, Chinnery PF (2020) Extreme heterogeneity of human mitochondrial DNA from organelles to populations. Nat Rev Genet 22: 106. https://doi.org/10.1038/s41576-02000284-x 2. Kukat C, Wurm CA, Spahr H, Falkenberg M, Larsson NG, Jakobs S (2011) Super-resolution microscopy reveals that mammalian mitochondrial nucleoids have a uniform size and frequently contain a single copy of mtDNA. Proc Natl Acad Sci U S A 108(33): 13534–13539. https://doi.org/10.1073/ pnas.1109263108 3. Peter B, Falkenberg M (2020) TWINKLE and other human mitochondrial DNA helicases: structure, function and disease. Genes (Basel) 11(4):doi:10.3390/genes11040408

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e1001041. https://doi.org/10.1371/journal. pbio.1001041 16. Martell JD, Deerinck TJ, Sancak Y, Poulos TL, Mootha VK, Sosinsky GE, Ellisman MH, Ting AY (2012) Engineered ascorbate peroxidase as a genetically encoded reporter for electron microscopy. Nat Biotechnol 30(11):1143. https://doi.org/10.1038/nbt.2375 17. Lam SS, Martell JD, Kamer KJ, Deerinck TJ, Ellisman MH, Mootha VK, Ting AY (2015) Directed evolution of APEX2 for electron microscopy and proximity labeling. Nat Methods 12(1):51–54. https://doi.org/10.1038/ nmeth.3179 18. Han S, Udeshi ND, Deerinck TJ, Svinkina T, Ellisman MH, Carr SA, Ting AY (2017) Proximity biotinylation as a method for mapping proteins associated with mtDNA in living cells. Cell Chem Biol 24(3):404–414. https://doi.org/10.1016/j.chembiol.2017. 02.002 19. Wang LJ, Hsu T, Lin HL, Fu CY (2020) Drosophila MICOS knockdown impairs mitochondrial structure and function and promotes mitophagy in muscle tissue. Biol Open 9(12). https://doi.org/10.1242/bio.054262 20. Morgenstern JP, Land H (1990) Advanced mammalian gene transfer: high titre retroviral vectors with multiple drug selection markers and a complementary helper-free packaging cell line. Nucleic Acids Res 18(12): 3587–3596. https://doi.org/10.1093/nar/ 18.12.3587 21. Naviaux RK, Costanzi E, Haas M, Verma IM (1996) The pCL vector system: rapid production of helper-free, high-titer, recombinant retroviruses. J Virol 70(8):5701–5705. https:// doi.org/10.1128/JVI.70.8.5701-5705.1996 22. Prieto J, Leon M, Ponsoda X, Sendra R, Bort R, Ferrer-Lorente R, Raya A, LopezGarcia C, Torres J (2016) Early ERK1/2 activation promotes DRP1-dependent mitochondrial fission necessary for cell reprogramming. Nat Commun 7:11124. https://doi.org/10. 1038/ncomms11124

Part IV Mitochondrial DNA Replication and Repair

Chapter 14 In Vitro Assays of TWINKLE Function Jay P. Uhler, Ulrika Alexandersson, and Maria Falkenberg Abstract TWINKLE is an essential helicase that unwinds the duplex mitochondrial genome during DNA replication. In vitro assays using purified recombinant forms of the protein have been an instrumental tool for gaining mechanistic insights about TWINKLE and its function at the replication fork. Here we present methods to probe the helicase and ATPase activities of TWINKLE. For the helicase assay, TWINKLE is incubated with a radiolabeled oligonucleotide annealed to an M13mp18 single-stranded DNA template. TWINKLE will displace the oligonucleotide, which is then visualized by gel electrophoresis and autoradiography. To measure the ATPase activity of TWINKLE, a colorimetric assay is used, which quantifies the release of phosphate upon ATP hydrolysis by TWINKLE. Key words mtDNA, Replication, In vitro, Mitochondria, DNA helicase

1

Introduction Helicases are ubiquitous motor proteins that catalyze the NTP-dependent unwinding of double-stranded nucleic acids [1]. Human mitochondria contain several helicases involved in DNA and RNA transactions. Among these is the replicative helicase TWINKLE, a member of the SF4 superfamily of ring-shaped helicases that shares sequence homology with bacteriophage T7 gene protein 4 primase-helicase [2]. TWINKLE acts at the replication fork to separate the two strands of the circular mitochondrial genome during mitochondrial DNA (mtDNA) replication [3]. This creates single-stranded DNA that the mitochondrial DNA polymerase γ (POLγ) uses as a template to duplicate the genome [4]. Pathogenic mutations in TWINKLE can disrupt the progression of the replication fork, which in turn can lead to mtDNA depletion, mtDNA deletions, and/or duplications [2, 5, 6].

Jay P. Uhler and Ulrika Alexandersson contributed equally with all other contributors. Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Over 20 mutations in TWINKLE have so far been linked to mitochondrial disease [1]. Purified TWINKLE forms a homohexamer that requires a replication fork-like DNA structure to initiate unwinding. TWINKLE encircles and translocates along one of the DNA strands in the 5′–3′ direction while excluding the other strand from the central channel, thereby breaking the bonds between the base pairs [1]. Translocation and unwinding are powered by the intrinsic nucleoside-triphosphatase (NTPase) activity of TWINKLE. Tight coupling between NTP hydrolysis and translocation depends on coordinated interactions between the monomers. TWINKLE has also been shown to be stimulated by mitochondrial single-stranded binding protein mtSSBP1 and to stimulate DNA POLγ activity [3, 4]. In this chapter, we describe in vitro methods to specifically determine the helicase and NTP hydrolysis activities of purified recombinant TWINKLE. The methods are relatively simple to perform, robust, versatile, and require no sophisticated equipment or niche expertise. For the helicase assay, purified recombinant TWINKLE is incubated with a radiolabeled double-stranded substrate in the presence of ATP. Reactions are then separated by gel electrophoresis for visualization and quantification of singlestranded products. The assay can be adapted to more specifically examine loading, processivity, or directionality and can be used to study disease-causing mutations in TWINKLE [5–7]. For the NTP hydrolysis assay, we use an ATPase assay that detects the release of free phosphate from ATP. This assay is also suitable for studying disease-causing mutations in TWINKLE [5–7]. To perform the assay, purified recombinant TWINKLE is incubated with ATP. TWINKLE hydrolyzes the ATP to release free phosphate. The hydrolysis activity can then be calculated indirectly by measuring the phosphate release. The free phosphate will then react with malachite green and molybdate to form a complex that can be measured on a spectrophotometer or plate reader.

2

Materials Prepare all solutions with ultrapure water and molecular grade reagents. Unless indicated otherwise, store reagents as instructed by the manufacturer.

2.1 DNA Helicase Template Preparation

1. M13mp18 single-stranded DNA (BioLabs). 2. DNA oligonucleotide: 5′-tail helicase oligonucleotide: 5′-40[T] GTAAAAC GACGGCCAGTGCC-3′ (see Note 1). 3. 10× Annealing buffer: 1 M NaCl, 0.2 M Tris–HCl pH 7.5.

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4. Illustra Microspin G-25 column. 5. T4 polynucleotide kinase (T4 PNK) (New England Biolabs). 6. 10× T4 PNK buffer (New England Biolabs). 7. EasyTides® adenosine 5′-triphosphate [γ-32P] ([γ-32P] ATP) (3000 Ci/mmol). 2.2 DNA Helicase Assay

1. Protein dilution buffer: 20 mM Tris–HCl pH 7.5, 10% glycerol, 0.5 mM EDTA, 0.2 M NaCl, 1 mM DTT, and 0.1 mg/ mL bovine serum albumin (BSA). To prepare a 10 mL stock, mix together all components and bring the volume up to 10 mL. Aliquot to working stocks by dispensing the solution into 1 mL aliquots and store at -20 °C. 2. Helicase assay buffer: 20 mM Tris–HCl pH 7.5, 5 mM MgCl2, 0.1 mg/mL BSA, 4 mM DTT, and 3 mM ATP (see Note 2). 3. 6× Stop solution: 90 mM EDTA, 6% SDS, 30% glycerol, and 0.25% (w/v) bromophenol blue. Store aliquots at 4 °C and keep a working aliquot at room temperature. Preheat to 37 °C just before use to be sure that the SDS is not precipitated. 4. 100 mM ATP solution. 5. Purified recombinant TWINKLE [8].

2.3

ATPase Assay

1. ATPase activity buffer: 20 mM Tris–HCl pH 7.5, 5 mM MgCl2, 0.1 mg/mL BSA, 4 mM DTT, and 0.5 mM ATP (see Notes 2, 3, 4, and 5). 2. Malachite Green Phosphate Assay kit (BioAssay System).

2.4 Gel Electrophoresis and Visualization

1. 10× Tris–boric acid EDTA (TBE) buffer. 2. 1× TBE running buffer: 100 mL 10× TBE and 900 mL water. Prepare fresh. 3. 40% Acrylamide/bis-acrylamide solution 19:1. 4. 10% (w/v) ammonium persulfate (APS). 5. TEMED. 6. Galileo Bioscience Reflection 2010 Dual-Gel Vertical Electrophoresis System (see Note 6). 7. Whatman® 3MM filter paper. 8. Clingfilm. 9. Super RX-N Medicine X-ray film (Fuji). 10. Film cassette with an intensifying screen (Kodak). 11. X-ray film processor (Kodak X-OMAT 1000). 12. FLA-7000 Phosphorimaging system (Fujifilm) (see Note 7). 13. Spectrophotometer (600–660 nm) or a plate reader (see Note 8).

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Methods

3.1 Helicase Oligonucleotide Labeling

1. 5′-end label the helicase oligonucleotide by combining the following in order: 1 μL (5 pmol) of the oligonucleotide, 17.5 μL water, 2.5 μL of 1× T4 PNK buffer, 1 μL (10 U) of T4 PNK, and 2.5 μL (3000 Ci/mmol) of [γ-32P] ATP in a 1.5 mL reaction tube. 2. Incubate at 37 °C for 60 min. 3. Incubate at 65 °C for 20 min to inactivate T4 PNK. 4. Remove unincorporated nucleotides using an illustra Microspin G-25 column according to the manufacturer’s instructions. 5. Store the labeled oligonucleotide at -20 °C. It is possible to use an oligonucleotide that is fluorescently labeled instead (see Note 9).

3.2 Helicase Template Preparation

The helicase template consists of the M13mp18 ssDNA template strand annealed to the 5′-end labeled helicase oligonucleotide. The construction of the helicase template is illustrated in Fig. 1. The circular M13mp18 ssDNA is combined with the labeled helicase oligonucleotide. It is possible to construct other types of templates if desired (see Notes 10 and 11). 1. Prepare an annealing reaction in an Eppendorf tube with 2.5 pmol of labeled oligonucleotide (step 3.1) and 2.7 pmol of M13mp18 ssDNA. Add water to reach a final volume of 100 μL. 2. Place the tube in a heat block and heat the sample at 80 °C for 5 min, then add 20 μL of 10× annealing buffer. 3. Cool the sample from 80 °C to room temperature by turning off the heat block. 4. For storage, aliquot and keep at -20 °C. The template yield will be 2.5 pmol per reaction, assuming 100% successful annealing (see Notes 12 and 13). The template can be diluted to a suitable working concentration in water (10 nM is recommended) and stored at -20 °C. 5' Helicase primer 5'

*

+

M13mp18 ssDNA

Anneal

*

40-nt flap

Helicase template

Fig. 1 Schematic of the helicase template construction. M13mp18 ssDNA is annealed to a 60 nt oligonucleotide labeled on the 5′-end (asterisk) creating a double-stranded region of 20 bp and a 40 nt 5′-tail

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TWINKLE

-

0

5

10

15

30

60

Time (min)

*

* 1

2

3

4

5

6

7

Fig. 2 Helicase activity reaction. A time curve with a constant amount of TWINKLE (1500 fmol, hexamer). TWINKLE was incubated with radiolabeled substrate in the presence of ATP and reactions were separated by gel electrophoresis. Lane 1: control for unwound ssDNA at time 0 (heated to 95 ° C), lane 2: 0 min, lane 3: 5 min, lane 4: 10 min, lane 5: 15 min, lane 6: 30 min, and lane 7: 60 min

3.3 Helicase Activity Reaction

The helicase reaction is carried out using the annealed helicase template (see Note 18). TWINKLE will displace the oligonucleotide (Figs. 1 and 2). Details on protein expression and purification are beyond the scope of this chapter and can be found elsewhere (see Subheading 3.2). In brief, express his-tagged version of TWINKLE (lacking the mitochondrial targeting sequence) in insect cells using the baculovirus system and purify using affinity and ion exchange chromatography. Measure the concentration and dilute the protein to the desired stock concentrations (see below). The reaction is sensitive to salt. Aliquot the proteins (3–10 μL aliquots), freeze them in liquid nitrogen, and store them at -80 °C. Perform everything on ice (a metal rack is recommended) unless stated otherwise. Add proteins last and thaw just before use. Always discard unused aliquots once thawed. On average, keep TWINKLE at approximately 500 nM (calculated as a hexamer). Table 1 shows the setup for a typical 15 μL reaction. The helicase reaction is carried out in several steps. In brief, a master reaction mixture is prepared from which TWINKLE is withheld, and the reactions are then started by the addition of TWINKLE. 1. In the simplest case of a time course experiment, for example, the following seven reactions are needed: one control and 0, 5, 10, 15, 30, and 60 min time points. Label seven microcentrifuge tubes 1–7, add 3 μL of 6× stop buffer to each, and set aside at room temperature so that the SDS does not precipitate.

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Table 1 Helicase reaction: a typical setup for a 15 μL reaction Component

Final concentration

Per 15 μL reaction

Water

–

9.75 μL

1 M Tris–HCl pH 7.85

20 mM

0.2 μL

0.5 M MgCl2

5 mM

0.15 μL

100 mM DTT

4 mM

0.6 μL

10 mg/mL BSA

100 μg/mL

0.15 μL

100 mM ATP

3 mM

0.15 μL

10 nM template

0.7 nM

1 μL

500 nM TWINKLE

100 nM

3 μL

Protein dilution buffer

–

0 μL

2. In a separate tube, prepare a master reaction mixture for eight reactions (seven reactions plus one extra for pipetting errors) in the following order: 74.8 μL water, 2.4 μL 1 M Tris–HCl (pH 7.5), 1.2 μL 0.5 M MgCl2, 4.8 μL 100 mM DTT, 1.2 μL 10 mg/mL BSA, 3.6 μL 100 mM ATP, and 8 μL of 10 nM helicase template (total 12 μL/reaction). 3. Mix gently and leave on ice. 4. In a separate tube, add 21 μL of 500 nM TWINKLE and leave on ice. 5. Add the master mix reaction to the TWINKLE mix, and mix gently (the concentration of TWINKLE at this step will be 100 nM/reaction). Immediately remove 15 μL and transfer to tubes 1 and 2 (prepared with 3 μL stop buffer in step 1), then mix and set aside at room temperature. Tube 1 is a control for 100% unwound ssDNA. Denature by heating at 95 °C for 5 min, and then leave the tube on ice for 1 min just before loading the gel. Tube 2 is the “0” time point. Immediately place the remainder of the reaction in a heating block set at 37 ° C and start the timer. 6. At the 5 min time point, remove 15 μL from the reaction mixture and add to tube 3, which contains a 3 μL stop solution. Mix and set aside at room temperature. Repeat this procedure at the 10, 15, 30, and 60 min time points for tubes 4, 5, 6, and 7, respectively. Samples can be stored at room temperature at this point but should be loaded on the gel within 15 min (see Note 14).

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1. Prepare a 10% non-denaturing polyacrylamide gel: mix 6.25 mL 40% acrylamide/bis-acrylamide solution 19:1, 2.5 mL 10× TBE buffer, and 16.25 mL H2O (see Note 15). Add 150 μL 10% APS and 32 μL TEMED before casting the gel (20 cm × 10 cm). Immediately insert a comb with a medium well width (30 μL capacity per well). Allow the gel to set. 2. Place the gel in a running tank and fill it with 1× TBE buffer so that the gel is just covered. 3. Load 15 μL of sample per well. 4. Run the gel at room temperature at 200 V for 45 min, or until the bromophenol blue dye has run approximately two-thirds into the gel.

3.5

Visualization

1. Remove the gel from the tank, and allow the excess buffer to drain away. 2. Place the trimmed gel on top of two 3MM filter papers (pre-cut to be slightly larger than the gel), allowing the gel to stick onto the paper. Cover the gel with cling film.

A

1.50

B

20

Phosphate conc. (µM)

3. Transfer the gel (including the 3MM backing) to a cassette and expose to a phosphorimaging screen for 1 h, or up to overnight. Scan using a phosphorscanner and re-expose if necessary. Alternatively, expose to autoradiography film at -80 °C for 1 h to several days, depending on signal strength. Develop in an X-ray film processor or by hand. A representative result of a helicase assay time course is shown in Fig. 3. If reactions are suboptimal, it may be due to template quality, protein quality, NaCl, or sample storage issues (see Note 16).

15

Absorbance (620nm)

1.25 1.00 0.75 0.50

10

5

0.25 0.00

0

0

10

20

30

Phosphate conc. (µM)

40

50

0

500

1000

1500

2000

TWINKLE conc. (fmol)

Fig. 3 ATPase assay. (a) Malachite Green Kit was used and a standard curve was made from the provided reagents according to the manufacturer’s instructions. (b) A mix with ATP was incubated with increasing concentrations of TWINKLE at 37 °C, Malachite Green reagent was added and absorbance was measured at 620 nm

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Table 2 ATPase reaction: a typical setup for a 20 μL reaction

Component

Final concentration

Per 20 μL reaction

Water

–

15.3 μL

1 M Tris–HCl pH 7.5

20 mM

0.4 μL

0.5 M MgCl2

5 mM

0.2 μL

100 mM DTT

4 mM

0.8 μL

10 mg/mL BSA

100 μg/mL

0.2 μL

100 mM ATP

0.5 mM

0.1 μL

500 nM TWINKLE hexamer diluted as indicated with protein dilution buffer

Variable

3 μL

3.6

ATPase Assay

The ATPase assay reaction uses the same protein as the helicase assay (see Note 18). As with the helicase assay, the ATPase assay reaction is sensitive to salt (see Notes 2 and 17). Perform everything on ice (a metal rack is recommended) unless stated otherwise. Add TWINKLE last and thaw just before use. Table 2 shows a typical set-up for a 20 μL reaction. In practice, several reactions are needed for an experiment and the reactions are carried out in several steps, as described in the following points. In brief, a master reaction mixture without the proteins is prepared. Separately, the protein is diluted to the correct concentration and then mixed with the master mixture to start the reaction. In the simplest case of a concentration experiment, for example, the following seven reactions are needed: 0, 12.5, 25, 37.5, 50, 62.5, and 75 nM. 1. Make an ATPase activity buffer for eight reactions (seven reactions plus one extra for pipetting errors) in the following order: 122.4 μL water, 3.2 μL 1 M Tris–HCl pH 7.5, 1.6 μL 0.5 M MgCl2, 6.4 μL 100 mM DTT, 1.6 μL 10 mg/mL BSA, 0.8 μL 100 mM ATP (total 17 μL/reaction). 2. Mix and leave on ice. 3. Label seven microcentrifuge tubes 1–7 and place them on ice. In tube 1, add 3 μL of protein dilution buffer; in tube 2, add 2.5 μL protein dilution buffer and 0.5 μL of TWINKLE (500 nM); in tube 3, add 2 μL protein dilution buffer and 1 μL of TWINKLE (500 nM); in tube 4, add 1.5 μL protein dilution buffer and 1.5 μL of TWINKLE (500 nM); in tube 5, add 1 μL protein dilution buffer and 2 μL of TWINKLE (500 nM); in tube 6, add 0.5 μL protein dilution buffer and

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2.5 μL of TWINKLE (500 nM); in tube 7, add 0 μL protein dilution buffer and 3 μL of TWINKLE (500 nM). 4. Transfer 16 μL of the master reaction mixture to each tube containing the different amounts of protein, and mix gently. Immediately place the tubes in a 37 °C heating block and start the timer. 5. After 60 min, stop the reactions by adding 60 μL of H2O and 20 μL of malachite green to each tube. 3.7 Visualization and ATPase Activity Calculation

1. Incubate samples for 20 min at room temperature for color development before measuring the absorbance at 620 nm on a spectrophotometer. 2. Determine the quantity of phosphate released (fmol/sec) by performing a standard curve with free phosphate (Fig. 3a) according to the manufacturer’s instructions (see Note 19). A representative result of an ATPase assay using increasing amounts of TWINKLE is shown in Fig. 3b.

4

Notes 1. The sequence of the first 40 nt at the 5′-end is non-complementary to the M13mp18 single-stranded DNA and forms a 5′ single-stranded flap needed for TWINKLE unwinding activity. The minimum flap length required for TWINKLE helicase activity is 20 nt. The following 20 nt of the oligonucleotide anneals to position 6287–6306 in the M13mp18 single-stranded DNA. The optimal dsDNA length for efficient unwinding is 20–25 bp. 2. The end concentration of NaCl in the reaction should be between 40 and 80 mM for optimal activity. Normally, no extra salt is needed since it is added together with the proteins that are kept in high salt buffer and diluted in protein dilution buffer. 3. The ATP concentration can vary between 0.1 and 4 mM. 4. The ATP can be exchanged for any other NTPs or dNTPs. ATP or UTP is the optimal substrate for TWINKLE. 5. The ATPase activity can be measured with and without the presence of 100 fmol of M13mp18 ssDNA. DNA can have a mild stimulatory effect on ATPase activity. If the assay is used to study other helicases or ATPases, the stimulation by different cofactors can be extended. 6. It is not necessary to use the Galileo Bioscience Reflection 2010 Dual-Gel Vertical Electrophoresis System; other gel systems can be used as well.

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7. Either X-ray film or phosphorimaging can be used for visualization. It is not necessary to have both systems available in the lab. 8. Assays can be executed in tubes, cuvettes, or multi-well plates. 9. The oligonucleotide can be labeled with Alexa647 at the 5′-end and visualized by using a ChemiDoc MP imaging System (Bio-Rad). 10. We have successfully used a range of templates, including oligo-based linear templates, oligo-based minicircles, as well as the larger M13mp18-based templates described here. Also, consider whether to label the template or the product. TWINKLE needs a fork structure to unwind the template (see Fig. 1). 11. The helicase template can be kept at -20 °C for several weeks. The half-time of the radioactivity is about 2 weeks, so simply expose the gels for a longer time if the template is older. 12. Annealing efficiency can be checked by 10% non-denaturing PAGE. 13. If annealing is efficient, no ssDNA oligonucleotide band of 60 nt should be visible in the gel. If an ssDNA band of 60 nt is visible, add more M13mp18 ssDNA and repeat the annealing. 14. Samples should not be stored. The displaced oligonucleotide can reanneal if the reactions are stored at room temperature. 15. Excess volume of the 10% non-denaturing polyacrylamide gel mix can be prepared and kept at +4 °C for up to 6 months. APS and TEMED should be added just before use. 16. There are several points to consider if helicase reactions are suboptimal. The quality of the proteins used in the assays is critical. The protein should have >95% purity and be free from contamination from nucleases and phosphatases. 17. Salt in the reactions comes from the protein or protein dilution buffer. The working salt range for the assays is 40–80 mM NaCl. At 100 mM NaCl, the activity decreases noticeably. Also, note that for some types of experiments, for example, protein titration, it is necessary to correct for salt (see Note 2 for more details). 18. Both the helicase assay and ATPase assay can be used to analyze other helicases or ATPases in mitochondria such as RNA helicases. Change the templates accordingly. 19. Prepare phosphate standards as described in the protocol provided by BioAssay Systems. The useful detection range is 0.02–40 μM phosphate. The hydrolysis activity of TWINKLE can be calculated from the standard curve.

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Acknowledgments This work was supported by grants to MF from the Swedish Research Council, Swedish Cancer Foundation, European Research Council, the IngaBritt and Arne Lundberg Foundation, and the Knut and Alice Wallenberg Foundation. References 1. Peter B, Falkenberg M (2020) TWINKLE and other human mitochondrial DNA helicases: structure, function and disease. Genes (Basel) 11:408 2. Spelbrink JN, Li FY, Tiranti V, Nikali K, Yuan QP, Tariq M, Wanrooij S, Garrido N, Comi G, Morandi L, Santoro L, Toscano A, Fabrizi GM, Somer H, Croxen R, Beeson D, Poulton J, Suomalainen A, Jacobs HT, Zeviani M, Larsson C (2001) Human mitochondrial DNA deletions associated with mutations in the gene encoding twinkle, a phage T7 gene 4-like protein localized in mitochondria. Nat Genet 28:223–231 3. Korhonen JA, Gaspari M, Falkenberg M (2003) TWINKLE has 5′ -> 3’ DNA helicase activity and is specifically stimulated by mitochondrial single-stranded DNA-binding protein. J Biol Chem 278:48627–48632 4. Korhonen JA, Pham XH, Pellegrini M, Falkenberg M (2004) Reconstitution of a minimal mtDNA replisome in vitro. EMBO J 23:2423– 2429

5. Korhonen JA, Pande V, Holmlund T, Farge G, Pham XH, Nilsson L, Falkenberg M (2008) Structure-function defects of the TWINKLE linker region in progressive external ophthalmoplegia. J Mol Biol 377:691–705 6. Holmlund T, Farge G, Pande V, Korhonen J, Nilsson L, Falkenberg M (2009) Structurefunction defects of the twinkle amino-terminal region in progressive external ophthalmoplegia. Biochim Biophys Acta 1792:132–139 7. Jemt E, Farge G, Backstrom S, Holmlund T, Gustafsson CM, Falkenberg M (2011) The mitochondrial DNA helicase TWINKLE can assemble on a closed circular template and support initiation of DNA synthesis. Nucleic Acids Res 39:9238–9249 8. Macao B, Uhler JP, Siibak T, Zhu X, Shi Y, Sheng W, Olsson M, Stewart JB, Gustafsson CM, Falkenberg M (2015) The exonuclease activity of DNA polymerase gamma is required for ligation during mitochondrial DNA replication. Nat Commun 6:7303

Chapter 15 Rolling Circle Replication and Bypass of Damaged Nucleotides Josefin M. E. Forslund, Gorazd Stojkovicˇ, and Sjoerd Wanrooij Abstract Faithful mitochondrial DNA (mtDNA) replication is critical for the proper function of the oxidative phosphorylation system. Problems with mtDNA maintenance, such as replication stalling upon encountering DNA damage, impair this vital function and can potentially lead to disease. An in vitro reconstituted mtDNA replication system can be used to investigate how the mtDNA replisome deals with, for example, oxidatively or UV-damaged DNA. In this chapter, we provide a detailed protocol on how to study the bypass of different types of DNA damage using a rolling circle replication assay. The assay takes advantage of purified recombinant proteins and can be adapted to the examination of various aspects of mtDNA maintenance. Key words Mitochondrial DNA, DNA replication, DNA polymerase, Rolling circle, DNA damage

1

Introduction Mitochondrial DNA (mtDNA) encodes for the central subunits of the oxidative phosphorylation system (OXPHOS), which is responsible for energy production in the cell. The integrity of mtDNA is therefore directly linked to proper cell function and is essential for the health of the organism. Correct transmission of the genetic information demands complete duplication of the DNA by the replication machinery. The mtDNA molecule is, however, susceptible to damage caused by endogenous and exogenous factors, which obstruct the polymerase progress and can lead to replication stalling [1–3]. When not resolved, replication fork stalling can lead to information loss and disease. Studies focusing on nuclear DNA have uncovered many strategies that cells use to prevent these DNA replication impediments from leading to the loss of genetic information [4]. On the contrary, there is still limited understanding of the fate of the mtDNA replisome when it encounters DNA lesions.

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Unlike the nucleus, the mitochondrion has only one replicative DNA polymerase, polymerase γ (POLγ), which is responsible for the synthesis of both strands of the circular mtDNA molecule. The POLγ holoenzyme consists of one catalytic subunit, POLγ A, and a dimer of the accessory subunit POLγ B [5]. POLγ, together with the replicative helicase TWINKLE [6] and the mitochondrial single-stranded DNA-binding protein (mtSSB) [7], forms the core mtDNA replisome. The POLγ holoenzyme synthesizes the new DNA strand and is stimulated by mtSSB, which suppresses secondary structures in the template. The TWINKLE helicase unwinds the upstream double-stranded DNA, providing the template strand for the polymerase. The ability of POLγ to bypass a variety of lesions, such as 8-oxo-G [8], UV-adducts [9], and ribonucleotides [10, 11], has been studied previously. However, on short linear substrates, the study of POLγ polymerase activity is somewhat restricted by the length of the substrate, which also impedes the efficient loading of mtDNA replisome partners. Instead, a circular substrate can be used to set up a rolling circle replication assay that addresses these shortcomings. By combining purified proteins and an artificial mini-circular template, we can reconstitute the synthesis of leading and lagging DNA strands in vitro [12]. Once initiated, leadingstrand DNA synthesis coupled with a continuous unwinding of the double-stranded template could, in principle, proceed indefinitely. With the ability to design the template DNA sequence, we can introduce specific types of DNA lesions and monitor their effects on the mtDNA replication machinery. This setup can also be used to test how other factors affect the bypass of damaged DNA [13, 14]. This chapter describes a detailed protocol of the in vitro rolling circle replication assay using the mtDNA replisome. As an example, replication of the DNA templates containing oxidatively damaged nucleotides is presented. The main steps are the preparation of a minicircle substrate, rolling circle replication assay, sequencing gel electrophoresis, and visualization of the reaction products.

2

Materials Use molecular grade reagents and ultrapure water. Reagents should be stored and handled according to manufacturers’ instructions.

2.1 Preparation of DNA Template 2.1.1 Circularization of Template and Purification

For the rolling circle assay, a linear 70-mer oligonucleotide is designed with either a non-damaged or a damaged base placed 30 nt from the 3′-end. A normal deoxyguanosine nucleotide base is used as the control template (non-damaged). For the damagecontaining template, either 8-oxo-7,8-dihydroguanine (8-oxo-G)

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Table 1 Example of oligonucleotides Name

Description

Sequence (5′–3′)

70 nt non-damaged

Normal deoxyguanosine nucleotide base

GAGGGGTATGTGATGGGAGGGCTA GGATATGAGGTGAGTTGAGTGGAGT TGGAAGTAGGCATACCCCTAT

70 nt 8-oxo-G

8-oxo-7,8-dihydroguanine

GAGGGGTATGTGATGGGAGGGCTA GGATATGAGGTGAGTT8oxoGAGTGGA GTTGGAAGTAGGCATACCCCTAT

70 nt AP

Abasic site

GAGGGGTATGTGATGGGAGGGCTA GGATATGAGGTGAGTTAPAGTGGAG TTGGAAGTAGGCATACCCCTAT

28 nt

Bridge oligo for template circularization

AGGCATACCCCTATGAGTTGGAAGT AGG

40 T28 (68 nt)

Primer for extension assay with 40 mer T-stretch overhang

T40CTCATAGGGGTATGCCTACTTCCAACTC

or abasic site (AP) are used here as examples (Table 1), but other types of DNA damage can also be included. To convert the linear oligo into a circular molecule, a 28-mer bridge oligonucleotide with 14 complementary bases at both ends of the 70-mer oligonucleotide is used. The 28-mer oligonucleotide promotes covalent circularization of the 70-mer oligonucleotide (Fig. 1). 1. Sodium chloride (NaCl). 2. Milli-Q water (mQ). 3. Heat block. 4. T4 DNA ligase (New England Biolabs). 5. 10× T4 ligase buffer (New England Biolabs): 500 mM Tris– HCl pH 7.5, 100 mM MgCl2, 10 mM ATP, and 100 mM DTT 6. Loading buffer: 95% formamide and, 250 mM EDTA. 7. Thin-layer chromatography plate (TLC). 8. UV lamp (254 nm). 9. Extraction buffer: 100 mM Tris–HCl pH 8.0, 500 mM NaCl, and 5 mM EDTA. 10. 100% and 30% acetonitrile. 11. Sep-Pak® plus C18 columns or similar. 12. MiniVac sample concentrator evaporator or equivalent. 13. TE buffer: 10 mM Tris–HCl pH 8.0 and 1 mM EDTA.

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Fig. 1 Schematic of substrate preparation. (a) The 28 nt bridge oligonucleotide is annealed to the 70 nt template to produce a circle. (b) The gap of the 70 nt template is ligated and the monomeric circles are purified from a denaturing gel. The monomeric circles will run slower than the unligated oligonucleotides, which will run at the same height as the linear control. (c) The purified circular template is annealed to a 68 nt long primer to produce the DNA substrate. Only 28 nt of the primer will be complementary to the template and a 40 nt flap will be formed. X indicates the position of either a normal dNMP or a damaged base

14. NanoDrop® equivalent.

ND-1000

UV-vis

spectrophotometer

or

Also, see Subheading 2.4. 2.1.2 DNA Substrate Preparation

1. 40 T28 primer for annealing to a circular template (Table 1, also see Note 1). 2. T4 polynucleotide kinase (Thermo Scientific). 3. 10× reaction buffer A (for forward reaction, Thermo Scientific): 500 mM Tris–HCl pH 7.6, 100 mM MgCl2, 50 mM DTT, and 1 mM spermidine.

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4. 4 μL γ32P ATP (3000 Ci/mmol 5 mCi/mL, and 20 μCi). 5. Illustra™ MicroSpin™ G-25 columns. 2.2 Purified Recombinant Proteins

Details on protein expression and purification are beyond the scope of this chapter and can be found elsewhere [13, 15] and only a brief description will be given for each protein (see Notes 2–5). All proteins lack the N-terminal mitochondrial targeting sequence. After purification, dilute proteins to the desired concentration (see Note 6), aliquot into small aliquots (2–5 μL), and snap-freeze in liquid nitrogen. Store the protein aliquots at -80 °C until use, and never re-freeze after thawing. 1. POLγ A (see Note 2). 2. POLγ B (see Note 3). 3. TWINKLE (see Note 4). 4. Mitochondrial single-stranded DNA-binding protein (mtSSB) (see Note 5).

2.3 In Vitro Replication Assay

1. Protein dilution buffer: dilute all proteins to the desired concentrations in their individual storage buffers or in one common dilution buffer (see Note 6). 2. Reaction buffer: 25 mM Tris–HCl pH 7.6, 10 mM MgCl2, 1 mM DTT, and 100 μg/mL BSA. 3. dNTPs: individual 100 mM solutions of dATP, dGTP, dCTP, or dTTP. 4. 100 mM ATP. 5. Stop buffer: 5% SDS and 250 mM EDTA. 6. Loading buffer: 95% formamide and 20 mM EDTA.

2.4 Sequencing Gel Electrophoresis

1. Owl S3S sequencing gel system with power supply or equivalent. 2. 50 well 0.4 mm comb with flat-tooth or equivalent. 3. 10× Tris–borate EDTA (TBE) solution. 4. 40% Acrylamide/bis solution (19:1). 5. Ultrapure urea. 6. N,N,N′,N′-tetramethylethylenediamine (TEMED). 7. 10% APS (ammonium persulfate solution, 1 g dissolved in 10 mL mQ). 8. Whatman® 3MM filter paper. 9. Model 583 gel dryer (BioRad) or equivalent. 10. Film cassette with phosphor storage screen. 11. Intensifying screen.

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12. Typhoon 9400 system. 13. ImageJ software or equivalent imaging processing software.

3

Methods

3.1 DNA Template Preparation

The 70 nt template with or without a damaged base is made into a minicircle by annealing a bridge oligo, which allows for ligation of the 5′- and 3′-ends of the template (Fig. 1a–b) (see Note 7). A 68 nt primer is then annealed to the mini circle template with a 40 nt overhang (Fig. 1c).

3.1.1 Preparation of Minicircle Template

To make a circular template, the 5′- and the 3′-ends of the 70 nt oligo need to be ligated. The 5′-end of the oligo needs to be phosphorylated to make ligation possible. Either order a phosphorylated 70 nt oligo or phosphorylate the oligo using a kinase (e.g., T4 Polynucleotide kinase, see the manufacturer’s instructions for the protocol). 1. In a total volume of 50 μL, prepare a mix of 24 μM bridge oligonucleotide and 12 μM phosphorylated template oligonucleotide, in the presence of 100 mM NaCl. 2. Vortex gently and place the tube in a heat block at 90 °C. Switch off the heat block and let it cool down (over several hours) to 30 °C before removing the tube (Fig. 1a). 3. Add 7 μL of 10× T4 ligase buffer, 1 μL of T4 ligase, and 12 μL of mQ. Incubate overnight at 16 °C (Fig. 1b). 4. Add 0.5 μL of T4 ligase and incubate for 4 h at 16 °C.

3.1.2 Purification of Circular Template on a Polyacrylamide Gel

1. Mix the ligated circular template (~70.5 μL) with 70 μL loading buffer containing 95% formamide and 20 mM EDTA (final concentration: 47.5% formamide, 10 mM EDTA). Prepare a second sample containing the unligated linear 70 nt oligonucleotide in the same loading buffer, to serve as a negative control. 2. Prepare a 10% polyacrylamide gel containing 1× TBE and 8 M urea (see Note 8 and Subheading 3.3). 3. Heat samples for 5 min at 95 °C and load the whole sample onto the gel (see Note 9). Run the gel for 1 h 45 min at 200 V. 4. Remove the gel from the cassette and place it on a TLC plate. Briefly irradiate the plate with UV light (254 nm) and mark the location of DNA (see Note 10). 5. Cut out the piece of gel containing the monomeric circular template, and place the gel fragments into a 15 mL polypropylene tube. 6. Add 2 mL of extraction buffer and incubate overnight at 60 °C.

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7. The next day, collect and save the supernatant (which contains DNA). To increase the recovery of DNA, add another 2 mL of extraction buffer to the gel fragments, and incubate at 60 °C for an additional 2 h. 8. Prepare a Sep-Pak® plus C18 column by washing with 10 mL 100% acetonitrile, 20 mL mQ, and 10 mL extraction buffer. 9. Load all 4 mL of DNA (overnight and 2 h fractions) on the prepared Sep-Pak® plus C18 column and collect the flowthrough. 10. Reload the flow-through three times to improve the DNA binding efficiency. 11. Wash the column with 20 mL of mQ. 12. Elute the DNA by applying 1 mL of 30% acetonitrile. Repeat this step four more times, separately collecting each 1 mL fraction. Most of the DNA should be in the first fraction. 13. Spin the tubes for approximately 4 h in a vacuum dryer to remove the acetonitrile. 14. Dissolve the DNA pellet in 100 μL TE buffer and determine the DNA concentration by measuring the absorbance at 260 nm using a NanoDrop® ND-1000 UV-Vis Spectrophotometer or equivalent. Calculate the molarity of the template, dilute to the desired working concentration with TE buffer, and store at -20 °C. 3.1.3 Labeling of the Primer Oligonucleotide at the 5′-End

1. Mix 1 μL of 10 μM 40 T28 oligonucleotide primer with 3 μL T4 polynucleotide kinase 10× reaction buffer A (final concentration 1×), 4 μL γ32P ATP (10 mCi/m), 1 μL T4 polynucleotide kinase (Thermo Scientific), and mQ up to a total volume of 30 μL. 2. Incubate at 37 °C for 1 h. 3. Add 20 μL mQ and clean up the reaction using Illustra™ MicroSpin™ G-25 columns by following the manufacturer’s protocol. This will give 50 μL of 200 nM γ32P-labeled primer.

3.1.4 Primer Template Hybridization

1. To anneal the primer to the template, in a tube gently mix 32 P-labeled primer with purified 70 nt circular oligonucleotide to reach a final concentration of 100 nM and 150 nM, respectively, in the presence of 100 mM NaCl (Fig. 1c). 2. Place the tube in a heat block at 95 °C for 2 min. Switch off the heat block and let it cool to 30 °C before removing the tube. 3. Store the DNA substrate at -20 °C until use.

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Fig. 2 Rolling circle replication by mtDNA replisome on minicircle substrate. (a) Overview of the DNA replication reactions. POLγ initiates replication and extends the primer until X (= normal or damaged base) is reached (step 2). If X is a normal base, POLγ can bypass and extend the primer until it reaches the 5′-end of the annealed primer (step 3). If X corresponds to a damaged base, this can lead to POLγ stalling (e.g. at 8oxoG site) or complete blockage of replication (e.g. at AP site). If POLγ bypasses the damage, the primer will be extended to the 5′ primer end (step 3). When reaching the primer end, TWINKLE can load onto the 5′-end flap and unwind the template. POLγ can continue the replication synthesis and mtSSB will coat the growing

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The rolling circle replication reaction is carried out using the primed minicircle substrate (Figs. 1 and 2a). Together, the 5′ primer flap and the template form a DNA replication fork, while the 3′-end of the primer is available for DNA synthesis. On a non-damaged template, replication will continue around the circular template until it reaches the 5′-end of the annealed portion of the primer. POLγ alone cannot displace a longer stretch of the primer, and the replication will therefore stop. If the helicase TWINKLE is added to the reaction, it will unwind the doublestranded DNA, displacing the (now extended) primer, and allowing the reaction to continue around the circular template. The displaced single-stranded DNA will be covered by mtSSB. In theory, the replication could continue indefinitely. If a damaged site is present in the DNA template, the inhibition of DNA synthesis by the DNA polymerase will be observed as an accumulation of products of a specific size. In the case of lesions that the polymerase can bypass, albeit with lower efficiency, long products (corresponding to several replications of the circle) will still be observed. A prominent band should be observed on the gel whenever the replisome tries to replicate past the lesion in the template, that is, on every full circle replication. On the other hand, if the polymerase cannot bypass the lesion, the primer will not be extended further than to the damaged site in the template. 1. Calculate the required amounts of reagents, taking into account the different reaction conditions to be tested, as well as the number of time points. Each experimental condition requires 10 μL of the reaction mixture (see Note 11). Depending on the complexity of the experiment, set up the number of reactions needed as in the example in Table 2. Remember to include a marker, a negative, and a positive control, if applicable (see Note 12). 2. Keep all reagents on ice unless stated otherwise. Thaw proteins just before use (see Note 13) and add to the reaction mix last, unless stated otherwise. 3. Prepare all buffers needed to the desired stock concentrations, and keep on ice. 4. In a prechilled tube, prepare a master mix (all components except for proteins) for the desired number of reaction conditions and time points to be tested.

ä Fig. 2 (continued) single-stranded DNA flap (step 4). (b) Visualization of mtDNA replisome replication on minicircle substrate containing normal dGMP (ND, lanes 1–5) or oxidative damage (8oxoG lanes 6–10 or abasic site AP lanes 11–15). Molecular markers are given in base pairs. A time course was performed with four time points (2, 6, 18, and 60 min). Stalling at strand displacement is indicated with an arrow and replication stalling at oxidative damage is indicated with arrowheads

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Table 2 The components of the reaction and their final concentrations. As an example, volumes that need to be pipetted are shown for a 40 μL reaction Reaction component (suggested stock concentration) Final concentration Example 40 μL reaction 1 M Tris–HCl pH 7.6

25 mM

1 μL

1 M MgCl2

10 mM

0.4 μL

100 mM DTT

1 mM

0.4 μL

10 mg/mL BSA

100 μg/mL

0.4 μL

100 mM ATP

4 mM

1.6 μL

100 μM dNTPs

10 μM

4 μL

100 nM 32P-40 T28/70 minicircle substrate

5 nM

2 μL 28.52 μL

mQ 20 μM mtSSB (tetramer)

250 nM

0.5 μL

900 nM TWINKLE

12.5 nM

0.556 μL

2000 nM POLγ A

12.5 nM

0.25 μL

2000 nM POLγ B (dimer)

18.75 nM

0.375 μL 40 μL

Final volume

5. Dilute proteins if needed (see Note 6). 6. Add mtSSB for pre-incubation on the substrate and leave it on ice for 5–10 min. 7. Initiate the reactions by adding TWINKLE helicase and the DNA polymerase subunits (POLγ A and B), and mix them gently. 8. Incubate reactions at 37 °C for 1–60 min depending on the desired extension. When several reaction conditions are to be tested in the same experiment, start the individual reactions sequentially, using regular time intervals (see Note 14). At the desired time point, stop the reaction by removing 10 μL of the reaction mixture, and transferring it to a tube containing 1.1 μL 5% SDS 250 mM EDTA. Keep the stopped reaction tubes on ice until all reactions are finished. Thereafter, incubate the proteins for 15 min at 50 °C. 9. Mix the samples with 11.1 μL loading buffer containing 95% formamide, 20 mM EDTA (final concentration 47.5% formamide, 10 mM EDTA), and store at -20 °C until analyzing them on a sequencing gel.

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1. Assemble the sequencing gel plates with 0.4 mm spacers. 2. Prepare a polyacrylamide gel solution containing 1× TBE, 10% acrylamide/bis-acrylamide (19:1), and 8 M urea. Slightly warm up the solution to dissolve the urea. Degas and filter the solution by passing the solution through a 0.45 μm vacuum filter (see Note 15). 3. For every 10 mL of gel solution, add 60 μL 10% APS and 10 μL TEMED. Swirl the solution and immediately pour it between the sequencing gel plates. 4. Immediately insert a flat-tooth comb (0.4 mm) and allow the gel to polymerize for at least 2 h. 5. After polymerization, remove the comb and the bottom spacer. Assemble the gel in the Owl S3S sequencing gel system. Fill the chambers with 1× TBE buffer and pre-run the gel for 30 min at 60 W. 6. When pre-running is nearly done, heat the samples for 5 min at 95 °C. Load 7 μL of sample in each well. To resolve the reaction products over the gel, run at 60 W for 2 h – 2 h 20 min, depending on the desired separation. 7. Stop the run and disassemble the gel apparatus. Remove the gel from the glass plates and transfer it to a Whatman® 3MM filter paper (see Note 16). Dry the gel in a Model 583 gel dryer or equivalent at 80 °C for 2 h and expose to a phosphor storage screen. 8. Scan the screen using a Typhoon 9400 Phosphor imager or an equivalent biomolecular imager (Fig. 2b). 9. Use ImageJ software or equivalent image processing software to analyze the results.

4

Notes 1. The 40 nt T stretch of the primer forms a 5′ flap as it is not complementary to the minicircle template. The flap should be at least 30 nt long since this is the minimum length needed for TWINKLE loading. Additionally, the primer and the minicircle template form a replication fork, which is also required for efficient TWINKLE loading. 2. Express the recombinant catalytic subunit of human DNA polymerase γ (POLγ A) with a 6xHis tag at its C-terminus in baculovirus-infected Spodoptera frugiperda (Sf9) cells. Lyse cells in lysis buffer (25 mM Tris–HCl pH 8.0, 500 mM NaCl, 10% glycerol, 10 mM imidazole, 5 mM 2-mercaptoethanol, and 1× protease inhibitors) and ultracentrifuge at 32000 rpm for 1 h to collect the total protein extract. Purify recombinant

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POLγ A on Ni2+-NTA agarose followed by an anion exchange (Q) chromatography column and a size exclusion column (Superdex 200 Increase 10/300 GL). 3. Express the accessory subunit p55 (POLγ B) in baculovirusinfected Sf9 cells with a 6xHis tag at the C-terminus. Lyse cells and ultracentrifuge as described for POLγ A to collect total protein extract. Purify POLγ B on Ni2+-NTA agarose followed by a Heparin column, a cation exchange (SP) chromatography column, and a size exclusion column (Superdex 200 Increase 10/300 GL). Calculate the POLγ B concentration as a dimer. 4. Express the TWINKLE helicase in baculovirus-infected Sf9 cells with a 6xHis tag at the C-terminus. Add lysis buffer (50 mM Tris–HCl pH 8.0, 600 mM NaCl, 20% glycerol, 20 mM imidazole, 5 mM 2-mercaptoethanol, and 1× protease inhibitors), ultracentrifuge and collect the total protein extract. Purify TWINKLE by running it over Ni2+-NTA agarose and a Heparin column. Calculate the concentration of TWINKLE as a hexamer. 5. Express mtSSB without a tag in baculovirus-infected Sf9 cells. Lyse cells in 20 mM Tris–HCl pH 8.0 with 500 mM NaCl, 10 mM imidazole, 5 mM 2-mercaptoethanol, and protease inhibitors. Ultracentrifuge for 1 h at 35000 rpm to collect whole cell extract. Buffer exchange into 30 mM HEPESKOH pH 7.6, 1 mM DTT, 0.25 mM EDTA, and 0.01% NP-40 using PD-10 columns. Purify mtSSB over an Affi-Blue column followed by Heparin, cation exchange chromatography and anion exchange chromatography columns. Alternatively, a hydroxyapatite column could be used instead of the SP and Q columns. Calculate mtSSB concentration as a tetramer. 6. The DNA replication reaction is sensitive to salt and the salt concentration should therefore be kept constant for reproducibility. Take into account the amount of salt coming from the addition of each protein, adjusting the stock protein concentrations if needed. A single type of dilution buffer can also be used if suitable for protein stability and activity. Doing so makes it easier to keep track of exact salt concentration and pH. 7. If a linear substrate is used, POLγ will only copy DNA until the end of the template. Also, note that TWINKLE is not necessary when using a linear template since no double-stranded DNA will be encountered by the DNA polymerase. If the linear form of the 70 nt template is to be used, move directly to Subheading 3.1.3. 8. It is not necessary to cast a big sequencing gel. A regular mini gel (8.6 × 6.8 cm) can also be used. Use 6% acrylamide/bisacrylamide (19:1) gel.

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9. To load the whole sample, load in several wells or use tape to increase the well size. By using larger wells, a reduced amount of gel is cut and this might improve DNA yield. 10. If the TLC plate is covered with a transparent sheet of plastic, it can be reused. Remember that the UV has to come from the top, therefore the normal lab UV tables cannot be used. 11. For example, if one reaction requires four different time points, a total volume of 45 μL (10 μL per time point and 5 μL extra for pipetting loses) should be prepared for that specific reaction. 12. A marker can be included for easier estimation of the DNA reaction product sizes. A marker can be prepared by labeling various DNA oligomers of known length as described in Subheading 3.1.3. A negative control without any protein (DNA substrate control) should always be included in order to be able to visualize the starting point of the reaction, as well as for possible quantification of the replication efficiency later on. 13. It is important to keep the reactions on ice when preparing the tubes since recombinant purified proteins can be sensitive to higher temperatures. For example, the POLγ A subunit is especially sensitive to elevated temperatures in the absence of DNA and the POLγ B subunit [16]. 14. To be able to run several reactions simultaneously, a defined time interval can be used for starting and stopping the reactions. Keep track of which order the reactions were started in and stop them in the same order. As an example, an experiment with four different reactions mixes (named WT1, WT2, mutant1, and mutant2) and three-time points (5, 10, and 30 min) could be performed like this: Start: WT 1 at 0 min, start WT 2 at 30 s, start mutant 1 at 1 min, start mutant 2 at 1.5 min Stop (5 min): WT 1 at 5 min, stop WT 2 at 5+30 s, stop mutant 1 at 5+1 min, stop mutant 2 at 5+1.5 min Stop (10 min): WT 1 at 10 min, stop WT 2 at 10+30 s, stop mutant 1 at 10+1 min, stop mutant 2 at 10+1.5 min Stop (30 min): WT 1 at 30 min, stop WT 2 at 30+30 s, stop mutant 1 at 30+1 min, stop mutant 2 at 30+1.5 min. The time interval between two samples needs to be adjusted according to the complexity of the experiment and the reaction times chosen. 15. Removing oxygen from the solution will both avoid air bubbles when pouring the gel and slow down the polymerization rate. The filtering will also remove potential small crystals of urea.

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16. When disassembling the two glass plates and the gel, be careful not to tear the gel since it is thin and very fragile. Make sure that one of the glass plates is silanized before pouring the gel. The silanization will allow the glass to detach from the gel, which can be then transferred to a Whatman paper. Cover the gel with a plastic folie in order to protect the gel from damage and the gel dryer from contamination during the drying.

Acknowledgments We thank Andreas Berner for his technical input. We thank Prof. Peter Burgers for the experiments carried out in his laboratory (Washington University in St. Louis). This work was supported by the Knut and Alice Wallenberg Foundation, and the Swedish Research Council. References 1. Cooke MS, Evans MD, Dizdaroglu M, Lunec J (2003) Oxidative DNA damage: mechanisms, mutation, and disease. FASEB J 17(10): 1195–1214. https://doi.org/10.1096/fj.020752rev 2. Young MJ (2017) Off-target effects of drugs that disrupt human mitochondrial DNA maintenance. Front Mol Biosci 4:74. https://doi. org/10.3389/fmolb.2017.00074 3. Copeland WC (2012) Defects in mitochondrial DNA replication and human disease. Crit Rev Biochem Mol Biol 47(1):64–74. https://doi.org/10.3109/10409238.2011. 632763 4. Ciccia A, Elledge SJ (2010) The DNA damage response: making it safe to play with knives. Mol Cell 40(2):179–204. https://doi.org/ 10.1016/j.molcel.2010.09.019 5. Lim SE, Longley MJ, Copeland WC (1999) The mitochondrial p55 accessory subunit of human DNA polymerase gamma enhances DNA binding, promotes processive DNA synthesis, and confers N-ethylmaleimide resistance. J Biol Chem 274(53):38197–38203. https://doi.org/10.1074/jbc.274.53.38197 6. Spelbrink JN, Li FY, Tiranti V, Nikali K, Yuan QP, Tariq M, Wanrooij S, Garrido N, Comi G, Morandi L, Santoro L, Toscano A, Fabrizi GM, Somer H, Croxen R, Beeson D, Poulton J, Suomalainen A, Jacobs HT, Zeviani M, Larsson C (2001) Human mitochondrial DNA deletions associated with mutations in the gene encoding Twinkle, a phage T7 gene 4-like protein localized in mitochondria. Nat

Genet 28(3):223–231. https://doi.org/10. 1038/90058 7. Mignotte B, Barat M, Mounolou JC (1985) Characterization of a mitochondrial protein binding to single-stranded DNA. Nucleic Acids Res 13(5):1703–1716. https://doi. org/10.1093/nar/13.5.1703 8. Graziewicz MA, Longley MJ, Copeland WC (2006) DNA polymerase gamma in mitochondrial DNA replication and repair. Chem Rev 106(2):383–405. https://doi.org/10.1021/ cr040463d 9. Kasiviswanathan R, Gustafson MA, Copeland WC, Meyer JN (2012) Human mitochondrial DNA polymerase gamma exhibits potential for bypass and mutagenesis at UV-induced cyclobutane thymine dimers. J Biol Chem 287(12): 9222–9229. https://doi.org/10.1074/jbc. M111.306852 10. Kasiviswanathan R, Copeland WC (2011) Ribonucleotide discrimination and reverse transcription by the human mitochondrial DNA polymerase. J Biol Chem 286(36): 31490–31500. https://doi.org/10.1074/jbc. M111.252460 11. Forslund JME, Pfeiffer A, Stojkovic G, Wanrooij PH, Wanrooij S (2018) The presence of rNTPs decreases the speed of mitochondrial DNA replication. PLoS Genet 14(3): e1007315. https://doi.org/10.1371/journal. pgen.1007315 12. Wanrooij S, Fuste JM, Farge G, Shi Y, Gustafsson CM, Falkenberg M (2008) Human mitochondrial RNA polymerase primes lagging-

mtDNA Rolling Circle Replication In Vitro strand DNA synthesis in vitro. Proc Natl Acad Sci U S A 105(32):11122–11127. https://doi. org/10.1073/pnas.0805399105 13. Korhonen JA, Pham XH, Pellegrini M, Falkenberg M (2004) Reconstitution of a minimal mtDNA replisome in vitro. EMBO J 23(12): 2423–2429. https://doi.org/10.1038/sj. emboj.7600257 14. Stojkovic G, Makarova AV, Wanrooij PH, Forslund J, Burgers PM, Wanrooij S (2016) Oxidative DNA damage stalls the human mitochondrial replisome. Sci Rep 6:28942. https:// doi.org/10.1038/srep28942

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15. Korhonen JA, Gaspari M, Falkenberg M (2003) TWINKLE has 5′ -> 3’ DNA helicase activity and is specifically stimulated by mitochondrial single-stranded DNA-binding protein. J Biol Chem 278(49):48627–48632. https://doi.org/10.1074/jbc.M306981200 16. Chan SS, Longley MJ, Copeland WC (2005) The common A467T mutation in the human mitochondrial DNA polymerase (POLG) compromises catalytic efficiency and interaction with the accessory subunit. J Biol Chem 280(36):31341–31346. https://doi.org/10. 1074/jbc.M506762200

Chapter 16 Studying Mitochondrial Nucleic Acid Synthesis Utilizing Intact Isolated Mitochondria Jelena Misic and Dusanka Milenkovic Abstract Mitochondria are eukaryotic organelles of endosymbiotic origin that contain their own genetic material, mitochondrial DNA (mtDNA), and dedicated systems for mtDNA maintenance and expression. MtDNA molecules encode a limited number of proteins that are nevertheless all essential subunits of the mitochondrial oxidative phosphorylation system. Here, we describe protocols to monitor DNA and RNA synthesis in intact, isolated mitochondria. These in organello synthesis protocols are valuable techniques for studying the mechanisms and regulation of mtDNA maintenance and expression. Key words Mitochondria, mtDNA, mtDNA maintenance and expression, in organello replication and transcription, Radioactive labeling of nucleic acids

1

Introduction Mitochondrial dysfunction plays a critical role in human ageing and various disease states, consistent with the central role of mitochondria in maintaining cellular energy homeostasis by the process of oxidative phosphorylation (OXPHOS) [1, 2]. Even though the vast majority of mitochondrial proteins are nucleus-encoded and are post-translationally imported into mitochondria, mitochondrial function is strictly dependent on the correct maintenance and expression of the mitochondrial DNA (mtDNA), which encodes 13 essential subunits of the OXPHOS system in mammals. Maintenance and expression of the mitochondrial genome require multicomponent machineries that comprise almost 100 different nuclear-encoded proteins [3]. Numerous disease-causing mutations are mapped to nuclear genes that affect mtDNA replication and expression, emphasizing the importance of versatile yet incompletely understood mechanisms controlling mtDNA function [1, 2, 4, 5].

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Here we describe methods for in organello mitochondrial nucleic acid synthesis using intact mitochondria isolated from tissues or cultured cells [6–11]. mtDNA and mtRNA are labeled by incorporation of radioactive nucleotides, resolved by agarose gel electrophoresis, and visualized from the membrane by autoradiography.

2

Materials

2.1 In Organello Replication and Transcription Assays

1. Qubit protein BR assay. 2. Bradford reagent.

2.1.1 Mitochondrial Protein Content Quantification 2.1.2

Buffers

1. Incubation buffer: 25 mM sucrose, 75 mM sorbitol, 10 mM K2HPO4, 100 mM KCl, 0.05 mM EDTA, 5 mM MgCl2, 10 mM Tris–HCl pH 7.4, 10 mM glutamate, and 2.5 mM malate. Adjust pH to 7.2 using HCl. The buffer can be prepared in advance and stored at -20 °C. On the day of the experiment, add 1 mg/mL BSA (fatty acid-free) and 1 mM ADP. 2. Washing buffer: 10% glycerol, 0.15 mM MgCl2, 10 mM Tris– HCl pH 6.8. Buffer can be prepared in advance and stored at 4 °C.

2.2 In Organello Replication Assay

1. dNTP, [α-32P]-3000 Ci/mmol; 10 mCi/mL.

2.2.1 mtDNA Radioactive Labeling 2.2.2

DNA Isolation

1. Reagents for DNA isolation. The exact reagents are dependent upon the method of choice. DNA isolation using the Gentra Puregene tissue kit (Qiagen) or phenol/chloroform extraction works equally well in our hands. 2. GlycoBlue coprecipitant. 3. TE buffer: 10 mM Tris–HCl pH 7.4 and 1 mM EDTA.

2.2.3

Southern Blot

1. Agarose gel: 0.8% agarose gel is used for resolving radioactively labeled mtDNA. Weigh 0.96 g of agarose and boil in 120 mL of 0.5× TBE (50 mM Tris, 45 mM boric acid, and 0.5 mM EDTA) until the solution is clear. No addition of a nucleic acid

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gel stain is necessary. The size of the tray used for gel polymerization is 16 × 13 cm (L × W). 2. Gel running buffer: 0.5× TBE. 3. 0.2 M HCl. 4. Denaturation buffer: 1.5 M NaCl and 0.5 M NaOH. 5. Neutralization buffer: 0.5 M Tris–HCl pH 7.4 and 1.5 M NaCl. 6. 20× SSC (saline-sodium citrate): 3 M NaCl, 300 mM trisodium citrate, and pH adjusted to 7.0. 7. Positively charged nylon membrane (pore size 0.45 μm). 8. 3MM Whatman paper. 9. Stack of paper towels. 10. Transfer tray. 11. Glass plates. 2.3 In Organello Transcription Assay

1. UTP, [α-32P]-3000 Ci/mmol; 10 mCi/mL. 2. Non-radiolabeled UTP.

2.3.1 mtRNA Radioactive Labeling 2.3.2

RNA Isolation

1. Reagents for RNA isolation. Exact reagents are dependent on the method of choice (e.g., TRIzol/chloroform extraction). 2. GlycoBlue coprecipitant.

2.3.3

Northern Blot

1. Agarose/formaldehyde gel: radioactively labeled mtRNA is resolved on 0.8–1.2% agarose gel containing formaldehyde. For the 1.2% gel, melt 0.96 g agarose in 57.6 mL of nuclease-free water in a microwave with frequent agitation until the agarose is completely in solution. Cool the solution down briefly and add 8 mL of 10× NorthernMax MOPS-based running buffer and 14.4 mL of 37% formaldehyde under a ventilation hood. Stir gently to avoid creating bubbles, and pour gel. The size of the tray used for gel polymerization is 16 × 13 cm (L × W). 2. Gel running buffer: 1× NorthernMax MOPS-based running buffer. 3. Positively charged nylon membrane (0.45 μm pore size). 4. Transfer buffer: 20× SSC. 5. 3MM Whatman paper. 6. Stack of paper towels. 7. Transfer tray. 8. Glass plates.

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Methods

3.1 In Organello Replication and Transcription Assays

The method assumes that mitochondria have been isolated (from mouse tissue, fruit flies, or cultured cells) using your method of choice (Fig. 1) (see Note 1). 1. Quantify mitochondria protein content using a suitable method (e.g., Bradford assay or Qubit protein BR assay). 2. Spin down mitochondria in safe-lock Eppendorf tubes at 8000 g for 4 min at 4 °C (see Note 2). Starting material amount is 500 μg – 1 mg per sample, depending on the experimental requirements (see Notes 3 and 4). 3. Wash mitochondrial pellet by resuspension in 500 μL of cold incubation buffer containing BSA and ADP (see Notes 5 and 6). 4. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 5. Repeat the washing step once more and continue with in organello replication (see Subheading 3.2) or transcription (see Subheading 3.3) assay.

3.2 In Organello Replication Assay

3.2.1

32

P Labeling-Pulse

Resuspend mitochondria in 500 μL of prewarmed (at 37 °C) (see Notes 5, 6, and 7) ready-made incubation buffer containing BSA, ADP, and non-radioactive nucleotides. The incubation buffer should contain all nucleotides except the one used for labeling (for example, 50 μM dGTP, 50 μM dCTP, and 50 μM dTTP if labeling using α-32P dATP). 1. In a dedicated isotope lab, add 20 μCi of α-32P dATP to each sample (2 μL of a 10 mCi/mL solution). 2. Seal the tubes contamination.

with

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to

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3. Rotate for 2 h at 37 °C (see Note 7). 3.2.2

Chase

In case you are not performing the chase step, then skip this paragraph and go directly to the washing steps (see Subheading 3.2.3). 1. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 2. Wash the mitochondrial pellet by resuspension in 500 μL of cold incubation buffer containing BSA and ADP (see Notes 5 and 6). 3. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 4. Resuspend the mitochondrial pellet in 520 μL of incubation buffer containing BSA, ADP, and all four nonradioactive nucleotides at 50 μM each (see Notes 5 and 6).

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an

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mtDNA deleted fragment

7S DNA

w t m ut

an t

Pulse

mtRNA

tRNA

Pulse

Fig. 1 Schematic overview of the experimental procedure with representative autoradiographies. In organello experiments were performed with isolated wild-type (wt) and mtDNA maintenance mutant mitochondria (mutant). For in organello replication experiment (upper panel) restriction digestion (NcoI, single cutter) of newly synthesized mtDNA was performed after the synthesis. Full-length mtDNA, deleted fragments as well as prematurely terminated replication products (7S DNA) were visualized. In in organello transcription experiment (lower panel) smear of mitochondrial rRNAs and mRNAs and the cloud of short tRNAs at the bottom of the figure are visualized. The illustrations were created with BioRender.com

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5. Divide samples into two 260-μL aliquots; one sample for the pulse and one for the chase. 6. For the pulse sample, continue with the washing steps in the next subheading. 7. For the chase sample, incubate for 2 h at 37 °C (see Note 7). Afterward, proceed with the same washing steps. 3.2.3

Washing

1. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 2. Wash mitochondrial pellet in 500 μL of cold washing buffer. 3. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 4. Repeat the washing step one more time. 5. Resuspend mitochondria in 500 μL of cold washing buffer and save 40 μL for an input/loading control. Add 12 μL of 4× Laemmli buffer to this fraction and freeze or use immediately to separate on SDS-PAGE gel followed by western blot analysis (see Note 8). 6. Centrifuge remaining 460 μL at 8000 g for 4 min at 4 °C (see Note 2).

3.2.4 Mitochondrial Lysis and DNA Extraction

1. Proceed with mtDNA extraction using your method of choice, for example, using the Gentra Puregene tissue kit (Qiagen) or phenol–chloroform extraction. 2. To ensure mtDNA enrichment and visualization, add 1 μL GlycoBlue coprecipitant (15 mg/mL) to each sample during the mtDNA precipitation step. 3. Dissolve mtDNA in a maximum of 30 μL of DNA hydration solution (water or TE buffer) at 42 °C for 1 h. 4. Optionally, mtDNA can be linearized by restriction digestion to visualize large deletions or to avoid looking at different topological forms of mtDNA. For instance, SacI and NcoI restriction enzymes cut murine mtDNA at single sites, while BamHI and XhoI are single-cutter restriction enzymes for human mtDNA. Heating for 5 min at 93 °C can be used for releasing 7S DNA for its better visualization (Fig. 1) [8]. 5. Take half of the isolated mtDNA, add the DNA loading dye and load them onto the agarose gel. 6. Run the gel for 15 h at 30 V. 7. Remove the gel from the tank and incubate in 0.2 M HCl for 10 min. 8. Rinse gel in H2O for 5 min. 9. Incubate gel in denaturation buffer for 20 min. 10. Rinse gel in H2O twice for 5 min each. 11. Incubate gel in neutralization buffer for 20 min.

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12. Rinse gel in H2O twice for 5 min each. 13. Finally, incubate gel in 20× SSC for 20 min. 14. Activate positively charged nylon membrane by incubating it briefly in H2O followed by incubation in 20× SSC. 15. Set up the capillary transfer overnight. Make a Whatman paperbridge over a glass plate with the edges soaked into a tray containing 20× SSC buffer. Add two additional Whatman paper sheets previously soaked in 20× SSC buffer. Make sure you remove all the air bubbles present. Place the gel on the top of the Whatman papers followed by the activated nylon membrane. Carefully remove all the remaining bubbles. Add two more Whatman paper sheets previously soaked in 20× SSC buffer. Assemble a stack of paper towels, place the glass plate on top and add some equally distributed weight of approximately 0.5 kg on top of everything. Proceed with transfer overnight. 16. Disassemble, transfer, and rinse the membrane briefly in 2× SSC. Seal the air-dried membrane in cling film and expose it for the desired time. Visualize the signal from the membrane using autoradiography (Fig. 1). 3.3 In Organello Transcription Assay

3.3.1

32

P Labeling-Pulse

Resuspend mitochondria in 1 mL of prewarmed (at 37 °C) (see Notes 5, 6, and 7) ready-made incubation buffer containing BSA and ADP. 1. In a dedicated isotope lab, add 60 μCi of α-32P UTP to each sample (6 μL of a 10 mCi/mL solution). 2. Seal the tubes contamination.

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3. Rotate for 1.5 h at 37 °C (see Note 7). 4. If a chase is being performed, divide the samples into two aliquots (one half for the pulse sample and another half for the chase sample, see Subheading 3.2.2), otherwise, continue with the total amount. 5. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 6. Resuspend mitochondrial pellet in 500 μL of incubation buffer containing BSA, ADP, and freshly added non-radioactive UTP to a final concentration of 2 mM. 7. Seal the tubes contamination.

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8. Rotate for 10 min at 37 °C (see Note 7).

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In case you are not performing the chase step, skip this paragraph and go to the washing steps, Subheading 3.3.3. 1. Rotate samples for additional 2 h (or according to desired time points) at 37 °C (see Note 7). 2. Proceed with the washing steps (see Subheading 3.3.3).

3.3.3

Washing

1. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 2. Wash mitochondrial pellet in 500 μL of cold washing buffer. 3. Centrifuge samples at 8000 g for 4 min at 4 °C (see Note 2). 4. Repeat the washing (see steps 2–3) one more time. 5. Resuspend mitochondria in 500 μL of cold washing buffer and save 40 μL for an input/loading control. Add 12 μL of 4× Laemmli buffer to this fraction and freeze or use immediately to separate on SDS-PAGE gel followed by western blot analysis (see Note 8). 6. Centrifuge the remaining 460 μL at 8000 g for 4 min at 4 °C (see Note 2).

3.3.4 Mitochondrial Lysis and RNA Extraction

1. Extract mtRNA using your method of choice (e.g., TRIzol®/ chloroform extraction). 2. To ensure mtRNA enrichment and visualization, add 1 μL GlycoBlue coprecipitant (15 mg/mL) to each sample during the mtRNA precipitation step and precipitate overnight at 20 °C. 3. Dissolve the samples in a maximum of 10 μL of RNase-free water at 60 °C for 10 min. 4. Add the RNA loading dye and incubate samples according to the manufacturer’s instructions. 5. Load samples on the agarose/formaldehyde gel. 6. Run the gel at 90 V for several hours until the RNA loading dye reaches two-thirds of the way down the gel. 7. Remove the gel from the tank and rinse in H2O twice for 10 min. 8. Incubate gel in 20× SSC for 20 min. 9. Activate positively charged nylon membrane by incubating briefly in H2O followed by incubating in 20× SSC. 10. Set up the capillary transfer overnight (see Subheading 3.2.4, step 15). 11. Disassemble, transfer, and rinse the membrane briefly in 2× SSC. Air-dry membrane, seal in cling film, and expose for the desired time. Visualize the signal from the membrane using autoradiography (Fig. 1).

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Notes 1. The quality of the mitochondria is crucial for those experiments; therefore, try to work fast and make sure that you have intact, coupled mitochondria. 2. If in organello replication or transcription is performed with fruit fly mitochondria, all the centrifugation steps are performed at 5000 g for 4 min at 4 °C. 3. For the in organello replication assay, use 300 μg, 500 μg or 1 mg of mitochondria depending on your experimental requirements. The initial amount can be split into fractions for 7S DNA visualization, analysis with and without linearization, or for a pulse-chase experiment. If 300 μg is used, splitting the sample is possible, but the obtained autoradiographic signal is weak. 4. For the in organello transcription assay, use 500 μg of mitochondria or 1 mg if a pulse-chase experiment is to be performed. 5. Mitochondria tend to aggregate in this buffer, which can be mitigated by initially resuspending the pellet in a smaller volume and then adding the rest. 6. Use tips with the end cut off using a clean pair of scissors when resuspending the mitochondrial pellet to avoid damaging them. 7. If the in organello replication or transcription assays are performed with fruit fly mitochondria, the incubation temperature is 30 °C. 8. A protein sample after mitochondrial nucleic acid synthesis is taken to control for the mitochondrial amount used for in organello synthesis assays. The protein sample is resolved by SDS-PAGE and the western blot is decorated with an antibody specific to a mitochondrial protein unrelated to mitochondrial maintenance or gene expression, for example, VDAC.

References 1. Gustafsson CM, Falkenberg M, Larsson N-G (2016) Maintenance and expression of mammalian mitochondrial DNA. Annu Rev Biochem 85:133–160 2. Protasoni M, Zeviani M (2021) Mitochondrial structure and bioenergetics in normal and disease conditions. Int J Mol Sci 22:586 3. Pfanner N, Warscheid B, Wiedemann N (2019) Mitochondrial proteins: from biogenesis to functional networks. Nat Rev Mol Cell Biol 1:267–284

4. Viscomi C, Zeviani M (2017) MtDNAmaintenance defects: syndromes and genes. J Inherit Metab Dis 40:587–599 5. Falkenberg M (2018) Mitochondrial DNA replication in mammalian cells: overview of the pathway. Essays Biochem 62:287–296 6. Gensler S, Weber K, Schmitt WE et al (2001) Mechanism of mammalian mitochondrial DNA replication: import of mitochondrial transcription factor a into isolated

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mitochondria stimulates 7S DNA synthesis. Nucleic Acids Res 29:3657–3663 7. Enriquez JA, Ramos J, Perez-Martos A et al (1994) Highly efficient DNA synthesis in isolated mitochondria from rat liver. Nucleic Acids Res 22:1861–1865 8. Matic S, Jiang M, Nicholls TJ et al (2018) Mice lacking the mitochondrial exonuclease MGME1 accumulate mtDNA deletions without developing progeria. Nat Commun 9:1202 9. Ruzzenente B, Metodiev MD, Wredenberg A et al (2011) LRPPRC is necessary for

polyadenylation and coordination of translation of mitochondrial mRNAs. EMBO J 31: 443–456 10. Enriquez JA, Lo´pez-Pe´rez MJ, Montoya J (1991) Saturation of the processing of newly synthesized rRNA in isolated brain mitochondria. FEBS Lett 280:32–36 11. Bratic A, Clemente P, Calvo-Garrido J et al (2016) Mitochondrial polyadenylation is a one-step process required for mRNA integrity and tRNA maturation. PLoS Genet 12: e1006028

Chapter 17 Functional Assessment of Mitochondrial DNA Maintenance by Depletion and Repopulation Using 2’,3’-Dideoxycytidine in Cultured Cells Ga´bor Zsurka, Genevieve Trombly, Susanne Scho¨ler, Daniel Blei, and Wolfram S. Kunz Abstract The manipulation of mitochondrial DNA (mtDNA) copy number in cultured cells, using substances that interfere with DNA replication, is a useful tool to investigate various aspects of mtDNA maintenance. Here we describe the use of 2′,3′-dideoxycytidine (ddC) to induce a reversible reduction of mtDNA copy number in human primary fibroblasts and human embryonic kidney (HEK293) cells. Once the application of ddC is stopped, cells depleted for mtDNA attempt to recover normal mtDNA copy numbers. The dynamics of repopulation of mtDNA provide a valuable measure for the enzymatic activity of the mtDNA replication machinery. Key words Replication of mtDNA, mtDNA copy number, Nucleoside reverse transcriptase inhibitor (NRTI), Zalcitabine, DNA polymerase γ (POLG), Quantitative PCR

1

Introduction One of the key aspects of mitochondrial genetics is the multi-copy nature of mtDNA [1]. The average number of mtDNA molecules within single cells is dependent on cell type and varies between a few hundred copies in peripheral mononuclear cells to several thousands in skeletal muscle fibers. Mitochondrial DNA replication is not directly coupled to cell division [2] but is rather believed to be regulated by the metabolic needs of the cell [3]. Substances that inhibit mtDNA replication have been used since the early days of mitochondrial genetics to generate cell lines that fully or partially lack mtDNA. Ethidium bromide proved to be an efficient agent for inducing complete loss of mtDNA in various immortal cell types. These mtDNA-free cells, referred to as ρ0 cells, were then fused with enucleated cells or blood platelets to

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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allow a repopulation with heterologous mtDNA and, thus, to generate the so-called cybrid (cytoplasmic hybrid) cell lines [4]. Since ethidium bromide has several side effects, such as inhibition of transcription of mitochondrially-encoded genes [5], other substances with less severe influence on cellular metabolism were tested for their ability to deplete mtDNA. 2′,3′-Dideoxycytidine (ddC, zalcitabine), a nucleoside initially developed for the treatment of HIV infections, has been proven to be proficient at depleting mtDNA copy number in cultured cells [6, 7]. Although ddC is less powerful at inducing complete loss of mtDNA, its interference with cellular function is limited to mtDNA replication, which makes it the preferred substance for generating cells partially depleted for mtDNA. In intact cells, the active nucleotide ddCTP is generated in the cytoplasm via phosphorylation of ddC by nucleoside kinases and imported into mitochondria [7]. Due to the lack of the 3′ hydroxyl group on the deoxyribose, no further nucleotides can be added to the 3’ DNA end once ddCTP is incorporated into the nascent DNA strand, which leads to the abortion of replication. Such aborted ends might be repaired through excision of the blocking nucleotide [8, 9], which might reduce the chain termination effect of ddC. Since the number of newly synthesized mtDNA molecules is suppressed and non-circular intermediates of mtDNA replication can be efficiently degraded in wild-type cells [10], the pre-existing pool of circular mtDNA is diluted as cells divide, which leads to a severe decrease in the average mtDNA copy number per cell within a few days [6]. Similar to viral reverse transcriptases, the mitochondrial replicative DNA polymerase γ (POLG) shows relaxed discrimination between natural deoxycytidine triphosphate and dideoxycytidine triphosphate in comparison to nuclear DNA polymerases [11], which makes ddCTP a good selective inhibitor of viral and mitochondrial DNA replication. In this chapter, we describe the use of ddC to induce partial loss of mtDNA in cultured human cells and a quantitative PCR method to monitor the mtDNA copy number alterations. We provide examples of how a lack of efficient mtDNA repopulation after releasing the replication blockade can be applied to assess dysfunctional mtDNA maintenance [12–14].

2 2.1

Materials Cell Culture

1. Dulbecco’s Modified Eagle Medium (DMEM, 4.5 mg/mL glucose and 1 mM sodium pyruvate) supplemented with 10% fetal calf serum (FCS), 10 U/mL penicillin and 10 mg/mL streptomycin, and 50 μg/mL uridine (see Note 1). 2. Phosphate-buffered saline (PBS).

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Table 1 Sequences of used PCR primers Primer ID

Sequence

MT3922F

5′-GAACTAGTCTCAGGCTTCAACATCG-3′

MT4036R

5′-CTAGGAAGATTGTAGTGGTGAGGGTG-3′

MT16520F

5′-CATAAAGCCTAAATAGCCCACACG-3′

MT35R

5′-CCGTGAGTGGTTAATAGGGTGATA-3′

KirF

5′-GCGCAAAAGCCTCCTCATT-3′

KirR

5′-CCTTCCTTGGTTTGGTGGG-3′

3. Trypsin–EDTA solution (0.05% trypsin and 0.5 mM EDTA in Hank’s Balanced Salt Solution (HBSS)). Store at 4 °C. 4. 5 mg/mL uridine stock solution. Dissolve 250 mg uridine powder in 50 mL sterile MilliQ water and a sterile filter. Store at 4 °C for up to 3 months. 5. 100 mM 2′,3′-dideoxycytidine (ddC). Dissolve 21.1 mg ddC powder in 1 mL sterile MilliQ water. Store at -20 °C. 6. CO2 incubator, at 37 °C and 10% CO2 (see Note 2). 7. Six-well cell culture plates. 2.2

DNA Isolation

1. QIAamp DNA Mini Kit. 2. UV spectrophotometer for DNA concentration determination.

2.3

Quantitative PCR

1. AceQ qPCR SYBR Green Master Mix (2× concentrated, including PCR buffer, dNTPs, DNA polymerase, and SYBR Green). 2. Primer stock solutions in water, 25 pmol/μL. For primer sequences, refer to Table 1. 3. Quantitative PCR device (single fluorescence detection is sufficient). 4. 96-well qPCR plates. 5. Transparent adhesive seals for 96-well qPCR plates.

3 3.1

Methods ddC Treatment

1. If you plan to investigate a new type of cell line, perform pilot experiments to determine the minimal efficient concentration of ddC that is able to reduce mtDNA copy number to less than 5% of the original copy number within 7 days in wild-type cells. We have found that 20 μM ddC was suitable for primary human

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Fig. 1 Depletion and repopulation of mtDNA in HEK293 cells. (a) Depletion with different ddC concentrations. Black, no ddC; green, 40 μM ddC; blue, 80 μM ddC; gray, 160 μM ddC; red, 320 μM ddC. The mtDNA primers used are located in the D-loop. (b) Depletion and repopulation of mtDNA in HEK 293 cells treated with 320 μM ddC. Black, minor arc mtDNA primers; red, D-loop mtDNA primers. The arrow indicates the time point (4 days), when the ddC administration was stopped

fibroblasts. In contrast, about 400 μM ddC is required for wild-type HEK293 cells to obtain a sufficient depletion of mtDNA (Fig. 1). Note that the rate of mtDNA copy number loss is a function of the growth rate of dividing cells. If your

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cells have a doubling time significantly longer than HEK293 cells (24–48 h), allow longer depletion times. 2. On the day before starting the ddC treatment, plate a number of cells that are equivalent to 20–25% confluence. For example, seed 5 × 105 HEK293 cells in each well of a six-well plate. 3. On the next day, collect cells from one well for the initial time point (see Subheading 3.2). 4. Add appropriate volumes of 100 mM ddC stock solution into the growth medium of the remaining wells to achieve the desired final concentration. 5. Grow the cells in a humidified 10% CO2 atmosphere at 37 °C. 6. You can obtain a detailed depletion/repopulation curve by collecting samples every other day. In order to do this, harvest the cells by trypsinization (see Subheading 3.2). Count the cells and re-seed an amount required for 20–25% confluence in a new well of a six-well plate in a fresh ddC-containing medium. For the rest of the cells, proceed with DNA isolation (see Subheading 3.3). Even if no trypsinization is required due to the low confluence of the cells in a well, fresh ddC medium should be applied every other day. 7. After 12 days of ddC treatment, aspirate the growth medium and replace it with a medium lacking ddC. Continue growing cells and collecting samples the same way, and allow 14 days of recovery without ddC. 3.2 Harvesting Cells (Six-Well Plate Scale)

1. Remove the growth medium. 2. Wash cells with 500 μL of 1× PBS, and remove PBS again. 3. Add 200 μL of trypsin–EDTA solution to the well. 4. After a few minutes, inspect cells under the microscope to ensure that cells have detached, then stop the reaction by adding 800 μL of growth medium. 5. Resuspend cells and transfer cell suspension into a 1 mL microcentrifuge tube. If re-plating part of the cells, count cells at this step and replate accordingly before pelleting the cells. 6. Pellet cells by centrifugation at 3000 rpm for 10 min, remove medium, and store the pellet at -20 °C.

3.3 DNA Isolation and Determination of Concentration

Follow the protocol provided by the manufacturer of the DNA isolation kit. You will find below a protocol based on the QIAamp DNA Mini Kit tissue isolation protocol. 1. Resuspend the cell pellet in 180 μL buffer ATL. Add 20 μL of proteinase K, vortex, and incubate at 56 °C in a shaking platform at 450 rpm until the cell pellet is dissolved (approximately

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1–2 h). Vortexing occasionally during this incubation may help to dissolve the pellet faster. 2. Briefly centrifuge to remove drops from the inside of the lid. 3. Add 200 μL of buffer AL to the sample. Mix by pulse-vortexing for 15 s. 4. Incubate at 70 °C for 10 min. 5. Briefly centrifuge to remove drops from the inside of the lid. 6. Add 200 μL of ethanol (96–100%) to the sample, and mix again by pulse-vortexing for 15 s. After mixing, briefly centrifuge the tube to remove drops from the inside of the lid. 7. Apply the mixture to the QIAamp Mini spin column placed in a 2 mL collection tube. Centrifuge at 6000 g for 1 min. 8. Place the QIAamp Mini spin column in a clean 2 mL collection tube, and discard the tube containing the filtrate. 9. Add 500 μL of buffer AW1 to wash the column. Centrifuge at 6000 g for 1 min. 10. Place the QIAamp Mini spin column in a clean 2 mL collection tube, and discard the collection tube containing the filtrate. 11. Add 500 μL buffer AW2 to further wash the column. Centrifuge at 20,000 g for 3 min. 12. Place the QIAamp Mini spin column in a clean 2 mL collection tube, and discard the collection tube containing the filtrate. Centrifuge at 20,000 g for 1 min to remove residual wash buffer. 13. Place the QIAamp Mini spin column in a clean 1.5 mL microcentrifuge tube, and discard the collection tube containing the filtrate. Add 200 μL of buffer AE or distilled water. 14. Incubate at room temperature for 5 min, and then centrifuge at 6000 g for 1 min to elute the DNA. 15. Repeat steps 13 and 14 for a second elution. Combine the two elutions and vortex briefly. 16. Pipette 2 μL of the DNA solution onto the tray of a SimpliNano device or similar UV spectrophotometer to determine DNA concentration by measuring optical density. 3.4

Quantitative PCR

Quantification of average mtDNA copy number per nucleus is performed by quantitative PCR comparing the amplification of an mtDNA-specific product with a PCR product specific for a singlecopy nuclear gene, KCNJ10. The protocol described here uses SYBR Green detection and is performed in triplicates at two different DNA concentrations in 25 μL reactions (see Note 3). 1. Prepare 120 μL volumes of 2 ng/μL DNA in water for each DNA sample.

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2. Prepare 80 μL volumes of 1 ng/μL DNA in water from each DNA sample by diluting 40 μL of the 2 ng/μL DNA solution with 40 μL of water. 3. Prepare two master mixes, one with mtDNA-specific primers (MT3922F and MT4036R) and one with primers specific for the single-copy nuclear gene KCNJ10 (KirF and KirR) (see Note 4; see Table 1 for primer sequences). A single reaction contains 12.5 μL of AceQ qPCR SYBR Green Master MixMix, 0.3 μL of each primer (25 pmol/μL), and 1.9 μL of water. 4. Pipette 15 μL of the master mix to each well of a 96-well plate. For triplicates at two different DNA concentrations, you can use one row of a 96-well plate for a single sample. In this case, columns 1–6 are used for reactions with mtDNA-specific primers (MT3922F and MT4036R), and columns 7–12 are used for reactions with KCNJ10-specific primers (KirF and KirR). 5. Pipette 10 μL of the diluted DNA samples to the appropriate wells and mix by pipetting up and down. In order to exclude DNA contamination of the reaction mixtures, perform no-DNA controls, adding 10 μL of water to each master mix. 6. Seal the plate with a transparent adhesive film. 7. Insert the plate into a qPCR device and run the following program: 95 °C for 15 min; 45 cycles of 95 °C for 15 s and 62.5 °C for 60 s (see Note 5). 3.5

Data Analysis

1. Inspect the fluorescence intensity curves for each reaction using the operating software of the qPCR device (e.g., CFX Manager, version 3.1 for Bio-Rad qPCR devices). Pay special attention to the initial baseline and the plateau line. A significant reduction of the maximal intensity in single reactions might indicate unsuccessful amplification. 2. Export the table of fluorescence intensity data for further analysis with external software such as SigmaPlot. For this, use the setting “No Baseline Subtraction” (see Note 6). 3. Perform fitting of the data points to a sigmoidal curve for each sample using a 4-parameter Chapman–Richard growth function as implemented in SigmaPlot. The function is described by the formula y = a0 + a (1 – e-bt)c (see Note 7). 4. Calculate Cq values for each reaction as defined by t value at the inflection point of the Chapman–Richard growth function (at d2/dt2 [y(t)] = 0). This can be achieved by using the formula Cq = (ln c)/b (see Note 8). 5. Inspect Cq values for each reaction. Confirm that identical triplicates result in similar Cq values (standard deviation 15,000 g (see Note 3).

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Fig. 7 Modifying treatments for 2D-AGE. The abundance of different non-linear nucleic acids can hinder the interpretation of a 2D-AGE image, and various treatments can be employed to remove or modify and thus identify specific molecule types. (a) Mouse brain mtDNA digested with ClaI, separated over 2D-AGE and probed with a cytB probe for the fragment containing the non-coding region. (b) The same mtDNA as in (a) treated with S1 nuclease before the 2D-AGE. S1 nuclease degrades single-stranded nucleic acids and can be used to clear up the picture, but it also truncates or destroys replication intermediates with single-stranded patches. (c) The same mtDNA as in (a) treated with T7 endonuclease I, an enzyme that cleaves cruciform DNA structures, Holliday junctions and heteroduplex DNA. Longer treatments reduce the overall signal, as the enzyme also cuts nicked double-stranded DNA. For a schematic representation of the 2D-AGE panels with labelled intermediate types, as well as a map of the restriction and probe coordinates, see Fig. 3. (d) Human AccI-digested adult heart mtDNA separated by 2D-AGE and probed for the restriction fragment containing the ND2 gene. (e) The same sample as in (d), but with induced branch migration between the first and second dimensions. A large proportion of molecules from the x-spike, but also some from the y-arc, have resolved into linear DNA molecules, thus migrating similar to the 1n spot in the second dimension. (f) Schematic representation of the analyzed fragment. Molecules arising through branch migration are marked in red

2.2 Hypotonic Homogenization of Cultured Cells

1. 1–2 mL (wet pellet volume) of cultured, exponentially growing cells and appropriate culture medium. Typical amounts are five 15 cm plates of 80% confluent HEK293 cells or ca. ten 15 cm plates of 80% confluent HeLa cells or fibroblasts. (see Note 1). 2. Centrifuge accommodating 15 mL tubes.

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3. 10 × Hypotonic homogenization buffer (HHB): 400 mM Tris–HCl pH 7.4, 250 mM NaCl, 50 mM MgCl2, and 100 mM EDTA. 4. 1 M DTT and 200 μg/mL BSA: dissolve in water and store in 1 mL aliquots at -20 °C. 5. 1 × and 0.1 × HHB. Dilute freshly from the 10 × HHB buffer stock and add 1 mg/mL BSA and 1 mM DTT. 6. Dounce homogenizer with a tight-fitting pestle (see Note 2). 7. 15 mL centrifuge tubes tolerating max, rcf of >15,000 g. 2.3 Homogenization of Tissues

1. Fresh tissue samples of >1 mL volume. A typical starting amount is one mouse brain, one-fourth of a mouse liver, or one mouse heart. Isolation of mitochondria and mtDNA is also possible from frozen brain or heart samples, but not from frozen liver. 2. Tissue homogenization buffer (THB): 225 mM Mannitol, 75 mM Sucrose, 20 mM Tris–HCl pH 7.4, 10 mM EDTA. Store at 4 °C. Dissolve DTT at a concentration of 1 M and BSA at a concentration of 200 μg/mL in water and store in 1 mL aliquots at -20 °C. Supplement THB freshly with 1 mM DTT and 1 mg/mL BSA. 3. Fine scissors or scalpel blades. 4. 50–100 mL glass beakers. 5. Dounce homogenizer with loose and tight-fitting pestle (see Note 2). 6. 15 mL centrifuge tubes tolerating max. rcf of >15,000 g.

2.4 Mitochondrial Nucleic Acid Isolation

1. 2 mL and 15 mL centrifuge tubes tolerating max. rcf of >15,000 g. 2. Centrifuges accommodating 2 mL tubes and 15 mL tubes with centrifugation speeds of ≥15,000 g. 3. Ultracentrifuge with swing-out rotor and 10–15 mL open-top tubes. 4. Sucrose gradient solutions: 1.5 M and 1.0 M sucrose in 10 mM HEPES pH 7.4, 10 mM EDTA (see Note 4). Store at 4 °C. 5. Homogenization buffer (HB) as described in Subheading 2.1, but without the addition of BSA and DTT. 6. Proteinase K, e.g., 20 mg/mL. Store at -20 °C. 7. Lysis buffer: 75 mM NaCl, 50 mM EDTA, 1% (v/v) Triton X-100, 20 mM HEPES pH 7.8. 8. Phenol/chloroform 1:1 pH 6.7–7.0. Store at 4 °C. 9. Chloroform. 10. Isopropanol.

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11. 70% Ethanol. 12. 20 mM HEPES–NaOH pH 7.2. 13. Spectrophotometer for nucleic concentration measurements (e.g., nanodrop). 2.5 Total DNA Extraction for 2D-AGE

1. 2 mL microtubes and centrifuge capable of centrifugation speeds of ≥15,000 g. 2. Lysis buffer: 75 mM NaCl, 50 mM EDTA, 1% (v/v) Triton X-100, 20 mM HEPES pH 7.8. 3. Proteinase K, e.g., 20 mg/mL. Store at -20 °C. 4. Phenol/chloroform 1:1, pH 6.7–7.0. Store at 4 °C. 5. Chloroform. 6. Ethanol 96%, pre-cooled at -20 °C. 7. 3 M sodium acetate pH 5.3. 8. 70% Ethanol. 9. 20 mM HEPES pH 7.2. 10. Spectrophotometer for DNA concentration measurements (e.g., nanodrop).

2.6 DNA Digestion and Two-Dimensional Neutral Agarose Gel Electrophoresis

1. Restriction enzymes with 10 × restriction buffer (see Table 1 for commonly used enzymes). 2. 1.5 mL microtubes and centrifuge. 3. 37 °C waterbath or incubator. 4. Phenol/chloroform 1:1, pH 6.7–7.0. Store at 4 °C. 5. 10 × DNA loading dye: 180 mM Tris–HCl pH 7.5, 2 mM EDTA, 50% glycerol, 0.1% xylene cyanol, 0.2% bromophenol blue (see Note 5). 6. Agarose with high separation ability, e.g., UltraPure Agarose (Invitrogen). 7. 1 × TBE electrophoresis buffer: 89 mM Tris, 89 mM boric acid, 2 mM EDTA. This buffer can be made or bought as 10 × concentrate (890 mM Tris, 890 mM boric acid, 20 mM EDTA). 8. 10 mg/mL ethidium bromide. Dissolve ethidium bromide in H2O and store at -20 °C. 9. Horizontal electrophoresis chamber for ca. 12 × 14 cm gels. 10. Horizontal electrophoresis chamber for ca. 20 × 25 cm gels with buffer circulation ports. 11. Peristaltic pump and tubing for buffer recirculation. 12. Cold room or fridge to run the second dimension electrophoresis.

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13. Power supply (≥150 V). 14. UV table or imager. 15. Glass or plastic dishes fitting the smaller gel tray. 16. Shaker. 2.7 Southern Blotting and Detection

1. Glass or plastic dish with flat bottom fitting the larger gel. 2. Shaker. 3. Depurination solution: 0.2 M HCl. 4. Denaturation solution: 0.5 M NaOH, 1.5 M NaCl. 5. Filter paper sheets cut to 20 × 25 cm. 6. Paper towels. 7. 6 × SSC: 900 mM NaCl, 90 mM sodium citrate pH 7.0. 8. UV crosslinker or device to incubate the membrane at 80 °C (e.g., gel dryer). 9. Hybridization oven with large hybridization tubes. 10. Hybridization buffer: 250 mM NaPO4 pH 7.0, 7% SDS, 1 mM EDTA. For 1 L, dissolve 30 g NaH2PO4 and 70 g SDS in 800 mL distilled water. If the SDS does not dissolve, warm the buffer slightly. Adjust the pH with 5 M NaOH and add 2 mL of 500 mM EDTA, pH 8.0. Adjust volume to 1 L with distilled water (see Note 6). 11. DNA probe (usually a PCR product of ca. 500 bp length). 12. Random primed labeling kit. 13.

P α-dCTP, 3000 mCi/mmol (see Note 7).

32

14. Wash buffer: 1 × SSC, 0.1% SDS (150 mM NaCl, 15 mM sodium citrate, 0.1% SDS pH 7.0). Make from a 20 × SSC concentrate (3 M NaCl, 0.3 M sodium citrate pH 7.0) and a 20% SDS concentrate in water. 15. Cling film. 16. Exposure cassettes (20 × 25 cm). 17. Sensitive X-ray film and enhancer screens for detection of 32P signals (e.g., Kodak MS with a Kodak MS intensifying screen), alternatively phosphor storage screens such as Fuji Bas-IP can be used together with a phosphor imager. 2.8 Additional Treatments and Variations of 2D-AGE

1. S1 nuclease. 2. RNase H. 3. T7 endonuclease I. 4. E.coli topoisomerase IV with the respective buffers provided by the supplier.

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5. Branch migration buffer: 10 mM Tris–HCl pH 8.0, 0.1 M NaCl, 0.1 mM EDTA pH 8.0. 6. Incubator or water bath at 65 °C with capability of mild agitation.

3

Methods Use only one of the methods in Subheadings 3.1, 3.2 or 3.3 for mitochondrial isolation depending upon the starting material, followed by extraction of mitochondrial nucleic acids as in Subheading 3.4. Alternatively, if starting from whole cells, then use the method in Subheading 3.5.

3.1 Homogenization of Cultured Cells with Cytochalasin

1. Grow cells to a subconfluent stage under suitable growth conditions. Harvest cells and pellet by centrifugation for 5 min at 800 g (see Note 1). 2. Remove the supernatant and resuspend the cells in 10 mL fresh culture medium. Add cytochalasin B to a final concentration of 20 μg/mL and mix by inversion. Return the cell suspension to a cell culture dish and incubate in a CO2-incubator for 30–60 min. 3. Transfer cells to a centrifugation tube (see Note 8) and centrifuge at 800 g for 3 min to pellet the cells. Discard the supernatant. 4. Resuspend the cell pellet in 5–15 mL of cold HB with BSA and DTT. 5. Transfer the cell suspension to a pre-chilled Dounce homogenizer and disrupt the cells using 12–20 strokes with a tightfitting pestle. 6. Assess the percentage of broken cells by trypan blue exclusion staining (see Fig. 8 and Note 9). Mix 20 μL of cell suspension with 20 μL of trypan blue solution and spread on a microscope slide. Nuclei released from broken cells are usually round, defined blue, while intact cells remain colourless. DNA released from disintegrated nuclei gives a punctate blue background staining. 60–70% disrupted cells and few disintegrated nuclei are ideal. If the cell breakage is insufficient, homogenize further using several additional strokes and check again. 7. Transfer the homogenate obtained to a prechilled 15 mL centrifugation tube and continue with Subheading 3.4.

3.2 Hypotonic Homogenization of Cultured Cells

1. Grow cells to a subconfluent stage under suitable growth conditions. Harvest cells and pellet by centrifugation for 5 min at 800 g (see Note 1). Remove the supernatant.

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Fig. 8 Optimization of cell homogenization for mtDNA isolation. Microscopy of trypan blue-stained cells allows for the optimization of cell homogenization and thus the isolation of purer mitochondria and better quality mitochondrial nucleic acids. HEK293 cells homogenized with 5, 14 or 20 Dounce strokes (as indicated) and stained with trypan blue are shown. Intact cells are bright and rounded, while free nuclei are smaller, spherical and stain blue. Broken nuclei form irregular clumps of stained grainy material. For most cell types is is possible to achieve 50–70% cell breakage with relatively low nuclear destruction (middle picture)

2. Wash the cell pellet by resuspension in 5 mL of 0.1 × HHB and centrifuge immediately again. Remove the supernatant. 3. Resuspend the cells in 2 mL of 0.1× HHB and incubate for 6 min on ice to allow swelling. 4. Transfer the cell suspension into a pre-chilled Dounce homogenizer and disrupt the cells with ca. 10 strokes (see Note 10). 5. Transfer cell suspension into a 15 mL tube and immediately add 200 μL of 10 × HHB, then mix by inverting several times. Continue with method Subheading 3.4. 3.3 Homogenization of Tissues

1. Cut the tissue piece coarsely and wash two to three times in 20–30 mL THB to remove blood, each time removing the liquid (see Note 11). 2. Cut the tissue with fine scissors into small pieces or mince with a scalpel blade. Wash the tissue again several times with THB. 3. Transfer tissue pieces with 5–10 mL of THB into a Dounce homogenizer. Use a loose pestle with 4–5 strokes to disrupt the tissue pieces. Continue for 5–10 strokes with a tight-fitting pestle to break cells. Transfer the tissue homogenate into a centrifuge tube and continue immediately with Subheading 3.4.

3.4 Mitochondrial Nucleic Acid Isolation

1. Centrifuge the cell or tissue homogenate obtained from Subheadings 3.1, 3.2 or 3.3 at 800 g for 5 min at 4 °C. Transfer the supernatant into a new centrifugation tube. The pellet consists of nuclei and undisrupted cells and can be discarded. Repeat this centrifugation and transfer once.

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2. Centrifuge the supernatant at 15,000 g for 10 min at 4 °C. Mitochondria and other cellular organelles will form a small but visible beige pellet. 3. Discard the supernatant completely and resuspend the crude mitochondrial pellet in 1 mL HB by pipetting up and down. 4. Create a two-step gradient in an ultracentrifuge tube by first pipetting 5 mL of 1.5 M sucrose solution into the tube, then gently overlaying this with 5 mL of 1.0 M sucrose solution (see Note 12). 5. Layer the mitochondrial suspension on top of the sucrose gradient and centrifuge in a swing-out rotor at 50,000 g at 4 °C for 30–60 min. During the centrifugation, the mitochondria will form a visible layer between the two sucrose layers. 6. Remove the sample suspension and most of the upper sucrose layer. Change to a fresh 1 mL pipette tip and recover the mitochondria between the sucrose layers by sucking them into the pipette tip. Transfer into a 2 mL microtube. 7. Dilute the mitochondrial suspension with ca. 1 volume of HB without the addition of BSA or DTT. Add Proteinase K to a final concentration of 20 μg/mL and incubate for 15 min on ice. 8. Pellet mitochondria by centrifugation at 16,000 g at 4 °C for 5 min. 9. Remove the supernatant and lyse the pellet in 1 mL of lysis buffer. Add proteinase K to a final concentration of 30 μg/mL and incubate 20 min on ice. 10. Under a fume hood, add 1 mL of phenol:chloroform (1:1), shake vigorously for 2 min and centrifuge for 5 min at ≥15,000 g to separate the phases. Transfer the upper aqueous phase into a fresh 2 mL tube, leaving any possible interphase behind. 11. Re-extract the sample twice with 1 mL phenol:chloroform in a similar fashion, centrifuging and always transferring the upper aqueous phase. 12. Extract the aqueous phase once with 1 mL chloroform alone, again centrifuging for 5 min at ≥15,000 g and transferring the upper phase into a fresh 2 mL tube. 13. Add 1 mL of isopropanol and 20 μL of 5 M NaCl to precipitate mitochondrial nucleic acids. Incubate for ≥1 h at -20 °C. 14. Centrifuge at ≥15,000 g at 4 °C for 30 min. Remove the supernatant and add 1 mL of 70% EtOH. Centrifuge for 5 min at ≥15,000 g at 4 °C, remove the supernatant completely and let the small pellet air-dry in the open tube for 10–15 min until it is completely transparent.

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15. Dissolve the nucleic acid pellet in 50–100 μL of 20 mM HEPES–NaOH pH 7.2 and determine the concentration by photometrical absorption measurement at 260 nm (see Note 13). Store the nucleic acid extract at -20 °C. 3.5 Total DNA Isolation

1. Harvest subconfluent cells from one 10 cm plate by a suitable method (e.g., using trypsin) and pellet by centrifugation for 5 min at 1000 g. Wash the cell pellet with 1 mL of cold PBS and centrifuge again for 5 min at 1000 g. Remove the supernatant completely. 2. Lyse the cell pellet in 500 μL lysis buffer, pipetting up and down until all visible clumps have disintegrated. 3. Add proteinase K to a final concentration of 0.8 μg/μL and incubate at 37 °C for 4 h to overnight. 4. Add 50 μL of 5 M NaCl and 500 μL of phenol:chloroform (1: 1) and mix on a turning wheel or shaker for 30–60 min. Separate the phases by centrifugation for 5 min at >15,000 g and transfer the upper aqueous phase into a fresh microtube. Avoid the transfer of any interphase. 5. Repeat the addition of one volume of phenol:chloroform, mixing, centrifugation and transfer of the aqueous phase as in step 4 until no interphase is visible. 6. Remove phenol traces by the addition of 500 μL of chloroform, mixing, centrifugation and transfer of the aqueous phase as in step 4 above. 7. Add 1 mL of cold 96% ethanol and 50 μL of 3 M sodium acetate (pH 5.3) to the aqueous phase. Mix well and incubate for ≥2 h at -20 °C or 30 min at -80 °C. 8. Pellet the precipitated DNA by centrifugation for 20 min at ≥16,000 g at 4 °C. 9. Remove the supernatant and add 1 mL of 70% EtOH to the pellet. Centrifuge for 5 min at ≥16,000 g at 4 °C. 10. Remove the supernatant completely and let the pellet air-dry in the opened tube for 15–30 min until the pellet is completely transparent. 11. Resuspend the DNA in 100 μL of 20 mM HEPES pH 7.2 and determine the concentration by photometrical absorption measurement at 260 nm (see Notes 13 and 14). Store the nucleic acid extract at -20 °C.

3.6 DNA Digestion and Two-Dimensional Agarose Gel Electrophoresis

1. Digest 5 μg of mitochondrial nucleic acid extract or 10 μg of total DNA with the restriction enzyme of choice. For this, dilute the nucleic acid extract into 42 μL of water and add 5 μL of 10 × restriction enzyme buffer and 3 μL of restriction enzyme. If two restriction enzymes are to be combined, reduce

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the dilution volume to 40 μL and add 5 μL of 10 × buffer and 2.5 μL of each enzyme. Incubate for ≥5 h at 37 °C. 2. If desired, add a DNA-modifying enzyme into the restriction mix (see Subheading 3.8), vortex briefly and incubate at 37 °C for the time indicated in Table 1. 3. Add 50 μL of phenol:chloroform (1:1) to the digestion mix, vortex for 30 s and centrifuge for 5 min at ≥16,000 g. 4. Transfer the upper aqueous phase into a fresh tube, avoiding taking any of the lower organic phase. Add 5 μL of DNA loading dye and mix. 5. Cast a mid-sized agarose gel (ca. 12 × 15 cm) using 150 mL of agarose in 1× TBE without ethidium bromide or other intercalating DNA dyes. Use a percentage of agarose appropriate for the fragment size analyzed as indicated in Table 2 and a comb with ca. 1 cm tooth width. 6. Submerge the gel in the electrophoresis chamber in 1× TBE buffer and remove the comb. 7. Load a DNA size marker in the first lane and up to four samples digested as described in Subheading 3.5. Keep 1–2 lanes empty between each loaded sample. 8. Separate the samples with the voltage and for the time indicated in Table 2 (see Note 15). 9. Transfer the gel on its tray into a dish, cover with 1× TBE containing 2 μg/mL ethidium bromide and shake gently for 15–20 min. 10. Place the stained gel onto a UV table to visualize the DNA. If the DNA is stained strongly, the gel can remain on its tray, otherwise remove the tray to increase the visibility of the lanes.

Table 2 Estimated run times and recommended voltages Gel type

Fragment size

A

B

1. Dimension

2. Dimension

3.5–5 kb

0.4% agarose, 16 h 35 V Bromophenol blue just left the gel

0.95%, 250 V 7 h or 120 V 16 h, +4 °C with buffer recirculation Bromophenol blue ca. 2 cm from end of the gel

13–17 kb

0.28% agarose, 24 h 35 V Xylene cyanol migrated ca. 40% of gel distance

0.58%, 50 V 72 h, room temperature with buffer recirculation Xylene cyanol migrated ca. 50% of gel distance

The precise values might vary between different models of electrophoresis chambers and can be adjusted accordingly

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11. Use a ruler and a scalpel to cut out the lanes containing the samples. Trim the cut-out lanes to 10 cm length, cutting ca. 1 cm below the band of the linear fragment. 12. Move the trimmed agarose slices onto the tray for the second dimension, arranging them, as shown in Fig. 1e. Cool the gel tray with the gel slices at 4 °C. 13. Melt the agarose for the second dimension gel in 400 mL of 1 × TBE. When the agarose solution has cooled to 60 °C, add ethidium bromide to 1 μg/mL and cast the gel around the first dimension slices. 14. When the gel has solidified, transfer it into the electrophoresis chamber and submerse it in 1 × TBE containing 1 μg/mL ethidium bromide. Assemble the pump and tubing for buffer recirculation during the electrophoresis. 15. Perform the second-dimension electrophoresis at 4 °C until the linear fragment of interest is ca. 1 cm before the end of its separation distance. Recirculate the buffer during the electrophoresis (see Note 16). 3.7 Southern Blotting and Detection

1. Remove the gel from the electrophoresis chamber and let it slide off the gel tray into a dish. 2. Add sufficient depurination solution to cover the gel and shake gently for 15 min. Remove the liquid and replace with fresh depurination solution for another 15 min (see Note 17). 3. Remove the liquid again and rinse the gel once for 30 s with water. 4. Add denaturation solution and shake the gel for 20 min, then remove the liquid and repeat for another 20 min with fresh denaturation solution. Remove the liquid again. 5. Cut a piece of nylon membrane to the size of the gel, moisten it in water and place it onto the gel (see Note 18). Remove any air bubbles under the membrane by rolling with a cylindrical object, e.g., a glass pipette. 6. Cut two sheets of filter paper to the size of the gel, moisten them with water and place them on the membrane. Again, roll out any air bubbles. 7. Stack ca. 10 cm of paper towel onto the filter paper and place a 1 kg weight on top. Let the capillary transfer continue for ≥5 h. 8. Remove the paper towel and filter paper. Lift the membrane off and rinse it briefly with 6 × SSC buffer to neutralize the pH. 9. Crosslink the nucleic acids onto the membrane by either incubating them between sheets of filter paper for 2 h at 80 °C or by UV crosslinking using 1200 J/cm2. The crosslinked membrane can be stored between filter paper at room temperature until it is hybridized.

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10. Place the membrane into a hybridization tube in a hybridisation oven preheated to 65 °C and rotate with ca. 20 mL preheated hybridization buffer for ≥30 min. 11. Label a probe detecting a part of the mtDNA fragment of choice by random primed labeling using a kit or published labeling methods [6] and remove unincorporated nucleotides by size exclusion chromatography (see Note 19). 12. Reduce the hybridization buffer amount in the hybridization tube to 5–10 mL. Denature the probe for 5 min at 95 °C and add it immediately to the hybridization liquid (see Note 20). 13. Incubate the membrane with the probe in hybridization buffer for ≥8 h. 14. Remove the probe liquid. Add 40 mL of wash buffer and incubate for 2–5 min before removing the wash buffer. Perform three additional wash steps, with 40 mL of wash buffer each time, for 20 min each. Collect the radioactive wash liquid and dispose of appropriately. 15. Take the membrane out of the hybridization tube and tap it dry between two sheets of filter paper. Wrap the membrane in cling film and expose it to a phosphor storage screen or to X-ray film sensitive to 32P radiation (see Note 21). Typical exposure times are 1–3 days for mtDNA and >7 days for total DNA samples. To compare the loading of the panels, also perform a short exposure of 1–2 h allowing estimation of the linear spot signal serving as a loading reference. 3.8 Additional Treatments and Variations of 2D-AGE

1. To identify single-stranded nucleic acids, perform two restriction digests of the same sample in parallel (Subheading 3.6, step 1).

3.8.1 Removal of SingleStranded Nucleic Acids

2. After digestion, add 50 U of S1 nuclease enzyme into the digest mix of one sample and incubate for 5 min at room temperature. 3. Immediately add 50 μL phenol:chloroform (1:1) to the reaction and mix by brief vortexing to stop the nuclease digestion. Centrifuge for 5 min at RT and transfer the upper aqueous phase into a fresh tube. Also extract the non-treated sample in a similar fashion. 4. Separate both samples in parallel on first and second dimension gels, blot and hybridize as described in Subheadings 3.6 and 3.7 above. 5. Compare the panels of non-treated and S1-treated samples. Any signal and feature visible in the non-treated, but not the treated sample consists of at least partially single-stranded molecules (see Fig. 7a, b). The digestion of ssDNA-containing molecules often creates smaller molecule species, that appear on the treated panel only.

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3.8.2 Removal of RNA from RNA:DNA Hybrids

1. To identify RNA:DNA hybrids, perform two identical digests of the same sample in parallel (Subheading 3.6, step 1). 2. After digestion, add to one sample 10 U of RNase H enzyme into the digest mix and incubate for 30 min at 37 °C. 3. Extract both samples with phenol:chloroform as described in Subheading 3.6, steps 3–4. Separate both samples in parallel on first and second dimensions, blot and hybridize as described in Subheadings 3.6 and 3.7 above. 4. Compare the panels of non-treated and RNase H-treated samples. Any signal and feature visible in the non-treated, but not the RNase H-treated sample, has extensive RNA stretches hybridized to DNA, as their removal changes the electrophoretic migration pattern.

3.8.3 Identification of Holliday Junctions and Other Branched Structures

1. Perform two identical digests of the same sample in parallel and separate them over the first dimension as described in Subheading 3.6, steps 1–8. Cut out both gel slabs. 2. Incubate one of the gel slabs in branch migration buffer for 4 h at 65 °C with gentle agitation. Keep the other gel slab in electrophoresis buffer at 4 °C. 3. After the incubation, place both gel slabs onto the large gel tray and continue with the second dimension as described in Subheading 3.6, steps 9–15. Southern blot membranes, as described in Subheading 3.7. 4. Compare the panels of non-treated and branch-migrated samples (see Fig. 7d, e). The treatment facilitates the movement of Holliday junctions and forks within a branched molecule, often leading to the branched structure falling apart into two separate linear double-stranded molecules that are visible on the linear arc. Y-shaped replication forks can reform into linear or x-shaped chicken-foot structures, again changing their migration in the second dimension.

3.8.4 Topological Analysis of mtDNA

1. Perform four different enzyme digests, as described in Subheading 3.6, step 1, using the same nucleic acid extract. Treat one sample with 10 U Topoisomerase IV in topoIV buffer for 30 min at 37 °C. Treat a second sample with 1 U T7 endonuclease, in its respective buffer, for 30 min at 37 °C (see Note 22). Digest the third sample with a restriction enzyme that cuts only once in mtDNA, e.g., XhoI for human or mouse mtDNA. Leave the fourth sample undigested. 2. Separate all four treatments on a first and second dimension of type B (see Table 2), but run the second dimension slightly longer, until the xylene cyanol dye has migrated ca. 50% of the panel distance. Continue with blotting and hybridization as described in Subheading 3.7.

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3. Compare the observed patterns in the differently treated panels. Spots present in the untreated sample but missing in the topoisomerase IV panel consists of supercoiled mtDNA, which are converted to relaxed forms by topoisomerase IV and gain in intensity. T7 endonuclease I removes various types of branched structures, such as replication and recombination intermediates. The treatment with a restriction enzyme reduces circular mtDNA forms and intensifies linear molecules, regardless of their topology.

4

Notes 1. Adherent cells can be harvested by trypsinization or any other gentle detachment. Suspension cells are directly pelleted by centrifugation. 2. While this protocol offers instruction for the use of a Dounce homogenizer, other homogenizer models can also be employed, such as Potter–Elvehjem-type homogenizers with teflon pestle. 3. If neccessary, the centrifugation steps to pellet mitochondria can be performed at 12,000 g, in which case the centrifugation time should be doubled. 4. When isolating mitochondria from brain tissue, replace the 1 M sucrose solution by a 0.8 M sucrose solution, as brain mitochondria have a slightly reduced density compared to mitochondria from cultured cells and other tissues. 5. Any regular DNA loading dye is suitable, but a high content of xylene cyanol and bromophenol blue is recommended to allow visualization of the migration during the run. 6. This buffer is similar to classical Church buffer, but with adjusted pH, reduced phosphate concentration and no BSA. Regular Church buffer is equally well suited. 7. Other probe labeling techniques, e.g., DIG- or fluorescence labeling, are possible but less sensitive and, therefore, do not allow the detection of weaker features. 8. Pipette the cell suspension slowly to avoid premature breakage of the cells at this step. 9. It is recommended to test the required homogenization strokes whenever a new cell line is used, as the fine-tuning of cell breakage allows isolation of more and higher quality mitochondrial nucleic acids. When using the same cell type routinely, this step can be omitted. 10. A microscopic check with trypan blue as described in Subheading 3.1, step 6, can be performed at this step to verify sufficient

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cell breakage, but should be conducted speedily, as the cells continue to swell during this time. 11. Most tissues have a higher density that THB, so that the tissue pieces sink quickly to the bottom of the container and the liquid can be decanted. Fat tissue floats, and the use of a sieve for the washes is advisable. 12. These volumes are for a 12 mL tube. For larger or smaller ultracentrifuge tubes, adjust the sucrose solution volumes to allow the overlaying with 1 mL of mitochondrial suspension. To achieve a clear front between the two sucrose solutions, tilt the tube while letting the upper sucrose solution run slowly along the tube wall onto the lower layer. 13. The sample contains both mitochondrial DNA and RNA, and an A260/A280 ratio of 2 or even higher is normal. 14. If the total DNA sample is hard to solubilize, it can be digested for 3–12 h with an enzyme not cutting the mtDNA in question, such as BglII for human and KpnI for mouse mtDNA. 15. If unsure whether the separation is sufficient, the marker lane can be cut off and stained with ethidium bromide. If needed, the electrophoresis can then be continued with the unstained rest of the gel. 16. While the direction of recirculation is not important, the pump speed should be sufficient to recirculate the chamber volume once every 20–30 min. Make sure that the recirculation is slow enough to avoid flushing the gel off the tray or fix the gel at the edges using tape barriers. Toward the end of the run, the separation distance of the gel can be checked on a UV table and the run continued if needed. When routinely using the same fragments, it is helpful to stab a few microliters of 10 × loading dye into the gel at the upper gel corner using a pipet tip, so that the migration of the resulting bromophenol blue spot allows a quick estimation of the already achieved separation distance. 17. As the large gel might become damaged when pouring the liquid out, it is recommended to use suction for solution changes. At the end of the depurination steps, the bromophenol blue spots in the gel should have changed colour from blue to yellow. This colour change is reversed again during the denaturation steps. 18. If the dish has a smooth bottom and is large enough for the whole gel then the next step can be performed with the gel remaining in the dish. Otherwise, cut the gel along the middle using a scalpel and transfer the two halves onto a large glass plate or a piece of cling film on the table. Do not attempt to transfer the gel as a whole as it might break. The membrane can

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be labeled at the edge with pencil to indicate the side carrying the nucleic acids as well as the experimental notes. 19. This removal of unincorporated nucleotides is not essential, but reduces the radiation exposure during the probe handling and also the background on the membrane. Suitable spin columns for this purpose are available, e.g., Mini Quick Spin Columns (Roche) or self-made Sephadex G50 columns of 0.5 mL volume. Alternatively, nucleotides can be removed by precipitation of the probe using 50 μL of 4 M NH4–acetate, 200 μL ethanol and 10 μg tRNA. Centrifuge for 5 min at 15,000 g, remove the supernatant, and dissolve the pellet in 100 μL H2O. 20. The more concentrated the probe is in the hybridization buffer, the stronger signal will be obtained. Therefore, the liquid amount should be just sufficient to flush over the membrane without drying during the hybridization. Avoid pipetting the probe directly onto the membrane, as this might cause artefacts. 21. If using a different form of labeling and detection, continue appropriately, e.g., for DIG-labeled probes continue with blocking, antibody incubation and luminescence detection. 22. Note that higher amounts of T7 endonuclease I also degrade non-branched mtDNA. Therefore, do not increase the enzyme amount or incubation time if the treatment decreases the level of, but fails to completely remove, a DNA form of interest. Restriction enzymes also often fail to cut circular mtDNA completely, especially in the ND5/6 gene region. The likely reason is that freshly replicated mtDNA contains patches of ribonucleotides or single-strandedness on the H-strand, blocking the restriction site.

Acknowledgements The authors would like to thank Ian Holt, Takehiro Yasukawa and Aurelio Reyes for establishing and sharing many 2D-AGE techniques for mitochondrial DNA. References 1. Belanger KG, Mirzayan C, Kreuzer HE, Alberts BM, Kreuzer KN (1996) Two-dimensional gel analysis of rolling circle replication in the presence and absence of bacteriophage T4 primase. Nucleic Acids Res 24: 2166–2175

2. Brewer BJ, Fangman WL (1987) The localization of replication origins on ARS plasmids in S. cerevisiae. Cell 51:463–471 3. Dandjinou AT, Larrivee M, Wellinger RE, Wellinger RJ (2006) Two-dimensional agarose gel analysis of DNA replication intermediates. Methods Mol Biol 313:193–208

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4. Gerhold JM, Aun A, Sedman T, Joers P, Sedman J (2010) Strand invasion structures in the inverted repeat of Candida albicans mitochondrial DNA reveal a role for homologous recombination in replication. Mol Cell 39:851–861 5. Goffart S, Cooper HM, Tyynismaa H, Wanrooij S, Suomalainen A, Spelbrink JN (2009) Twinkle mutations associated with autosomal dominant progressive external ophthalmoplegia lead to impaired helicase function and in vivo mtDNA replication stalling. Hum Mol Genet 18:328–340 6. Green MR, Sambrook J (2019) Random priming: labeling of purified DNA fragments by extension of random oligonucleotides. Cold Spring Harb Protoc 2019 7. Hyvarinen AK, Pohjoismaki JL, Reyes A, Wanrooij S, Yasukawa T, Karhunen PJ, Spelbrink JN, Holt IJ, Jacobs HT (2007) The mitochondrial transcription termination factor mTERF modulates replication pausing in human mitochondrial DNA. Nucleic Acids Res 35:6458–6474 8. Joers P, Jacobs HT (2013) Analysis of replication intermediates indicates that Drosophila melanogaster mitochondrial DNA replicates by a strand-coupled theta mechanism. PLoS One 8:e53249 9. Kunnimalaiyaan M, Shi F, Nielsen BL (1997) Analysis of the tobacco chloroplast DNA replication origin (oriB) downstream of the 23 S rRNA gene. J Mol Biol 268:273–283

10. Lewis SC, Joers P, Willcox S, Griffith JD, Jacobs HT, Hyman BC (2015) A rolling circle replication mechanism produces multimeric lariats of mitochondrial DNA in Caenorhabditis elegans. PLoS Genet 11:e1004985 11. Manchekar M, Scissum-Gunn K, Song D, Khazi F, McLean SL, Nielsen BL (2006) DNA recombination activity in soybean mitochondria. J Mol Biol 356:288–299 12. Mettrick KA, Weaver GM, Grainge I (2020) Neutral-neutral 2-dimensional agarose gel electrophoresis for visualization of E. coli DNA replication structures. Methods Mol Biol 2119:61–72 13. Pohjoismaki JL, Goffart S, Tyynismaa H, Willcox S, Ide T, Kang D, Suomalainen A, Karhunen PJ, Griffith JD, Holt IJ, Jacobs HT (2009) Human heart mitochondrial DNA is organized in complex catenated networks containing abundant four-way junctions and replication forks. J Biol Chem 284:21446–21457 14. Reyes A, Yang MY, Bowmaker M, Holt IJ (2005) Bidirectional replication initiates at sites throughout the mitochondrial genome of birds. J Biol Chem 280:3242–3250 15. Yasukawa T, Yang MY, Jacobs HT, Holt IJ (2005) A bidirectional origin of replication maps to the major noncoding region of human mitochondrial DNA. Mol Cell 18: 651–662

Chapter 19 Quantitative Analysis of Nucleoside Triphosphate Pools in Mouse Muscle Using Hydrophilic Interaction Liquid Chromatography Coupled with Tandem Mass Spectrometry Detection Sushma Sharma, Ziqing Kong, Shaodong Jia, Phong Tran, Anna Karin Nilsson, and Andrei Chabes Abstract Defects in deoxyribonucleoside triphosphate (dNTP) metabolism are associated with a number of mitochondrial DNA (mtDNA) depletion syndromes (MDS). These disorders affect the muscles, liver, and brain, and the concentrations of dNTPs in these tissues are already normally low and are, therefore, difficult to measure. Thus, information about the concentrations of dNTPs in tissues of healthy animals and animals with MDS are important for mechanistic studies of mtDNA replication, analysis of disease progression, and the development of therapeutic interventions. Here, we present a sensitive method for the simultaneous analysis of all four dNTPs as well as all four ribonucleoside triphosphates (NTPs) in mouse muscles using hydrophilic interaction liquid chromatography coupled with triple quadrupole mass spectrometry. The simultaneous detection of NTPs allows them to be used as internal standards for the normalization of dNTP concentrations. The method can be applied for measuring dNTP and NTP pools in other tissues and organisms. Key words Deoxyribonucleoside triphosphates, Liquid chromatography, Triple quadrupole mass spectrometry, ZIC–HILIC, Differentiated tissues

1

Introduction Deoxyribonucleoside triphosphates (dNTPs) are the building blocks of DNA, and changes in their concentrations affect DNA replication fidelity and genome stability [1–4]. Alterations in dNTP pools have been extensively studied in relation to multiple biochemical processes [5–7]. In actively proliferating eukaryotic cells, the bulk of the de novo dNTP synthesis occurs in the cytosol from where dNTPs are transported to the nucleus and mitochondria. Non-dividing cells rely

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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heavily on salvage pathways, although de novo dNTP synthesis in these cells remains essential. Recent studies have shown that changes in dNTP pools that arise in the cytosol of Saccharomyces cerevisiae or mouse cells due to mutations in dNTP-metabolizing enzymes are directly transmitted into the mitochondria [8, 9]. This conclusion is based on the correlation between the overall cellular dNTP/nucleoside triphosphate (NTP) ratios and frequencies of ribonucleotide incorporation into mitochondrial DNA (mtDNA) (a method for detection of ribonucleotides in mtDNA is described in Chapter 21 of this volume), and this supports the notion of unregulated nucleoside and nucleotide transport across the mitochondrial membrane. These studies are in agreement with the proposed rapid interchange of mitochondrial and cytosolic nucleotide pools in cultured mammalian cells [10, 11]. Thus, changes in the overall dNTP pools in cells or tissues closely reflect changes in the intramitochondrial dNTP pools available for mtDNA replication. Several analytical methods have been developed to measure dNTPs and NTPs. The traditional DNA polymerase and radioactivity-based methods used for quantification of dNTP pools are sensitive, but they cannot simultaneously measure both dNTP and NTP concentrations [12–14]. Furthermore, higher concentrations of NTPs in the cells compared to the corresponding dNTPs can lead to the incorporation of NTPs into the DNA by DNA polymerases, thereby influencing the results [15–17]. Apart from the enzymatic assays, strong anion exchange highperformance liquid chromatography (HPLC) coupled with ultraviolet (UV) detection [18, 19] has also been used for measuring all eight canonical dNTPs and NTPs with high specificity and accuracy from biological samples containing high amounts of nucleotides [20, 21]. However, because dNTP pools in non-proliferating cells are very low, their quantification in differentiated tissue samples using HPLC–UV methods is challenging [22]. Methods based on HPLC coupled with mass spectrometry (MS) are more sensitive, but they are either not able to detect all NTPs [23] or require ion-pair reagents in the mobile phase, which can lead to the suppression of mass detection by affecting the performance of MS during ionization of the analytes and can contaminate the ion source [24]. Here, we describe a sensitive and ion-pairing-free method for the simultaneous separation of all eight dNTPs and NTPs in mouse skeletal muscle tissue extracts by utilizing a silica phosphorylcholine-based SeQuant® ZIC®–cHILIC column coupled with tandem MS. The method has shorter analysis times and easier sample cleanup steps prior to LC–MS. The steps involved in this method include cell lysis by homogenization and nucleotide extraction with trichloroacetic acid (TCA), Freon/trioctylamine neutralization of the extracts, boronate column separation of

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dNTPs from the nucleotide mixture, solid-phase extraction for sample cleaning and concentrating, and finally ultra-high-performance liquid chromatography (UHPLC) gradient separation coupled with MS detection of dNTPs and NTPs. This protocol, which is an adaptation of previously published methods [19, 25, 26], allows for the analysis of dNTP and NTP levels in differentiated mouse tissues, but can easily be adapted for other tissues and organisms.

2

Materials Solutions are prepared in ultrapure water from a Milli-Q water system using analytical-grade reagents unless indicated otherwise. No sodium azide is added to the solutions. All solutions are stored at room temperature unless indicated otherwise.

2.1 Collection of Muscle Tissue Sample

1. Cell lysis solution: 12% TCA (w/v) in 15 mM MgCl2. Weigh 3.0 g of TCA in a clean 50 mL conical tube and add 2.5 mL of 150 mM MgCl2 to the tube and make up the volume to 25 mL with sterile water. Store TCA powder and solution at 4 °C. Work with TCA in a fume hood. The prepared solution is stable for about 4 weeks. 2. 150 mM MgCl2: weigh 15.25 g of MgCl2·6H2O and dissolve in 450 mL water. Make up the volume to 500 mL and filter the solution through a 0.2 μm membrane filter into another clean glass bottle. The solution is chemically stable and can be stored at room temperature for several months. 3. Liquid nitrogen.

2.2 Extraction of dNTPs and NTPs from Muscle Tissue Sample

1. 3.2 mm stainless-steel beads.

2.3 Separation of dNTPs from NTPs with a Boronate Affinity Column

1. 0.1 M sodium boronate solution: dissolve 3.1 g of boric acid (H3BO3) in 480 mL of water in a beaker. Adjust the pH to 8.9 with 5 M sodium hydroxide. Transfer the solution to a graduated cylinder and make up to 500 mL with water. We usually do not filter this solution. The solution is chemically stable and can be stored at room temperature for several months.

2. Bead beater/homogenizer for beads (see Note 1). 3. Neutralization solution: 2.8 mL 98% trioctylamine, 10 mL Freon (1,1,2-trichloro-1,2,2-trifluoroethane), mix well (see Note 2).

2. Boronate affinity columns: hydrate the Bio-Rad Affi-Gel Boronate media overnight in boronate affinity column washing

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buffer and pack into the Bio-Rad Glass Econo-column. Store the packed columns at 4 °C (see Note 3). 3. Ammonium bicarbonate (ambic) buffer: 50 mM (NH4)2CO3, 15 mM MgCl2. Ambic buffer should be freshly made each time from stock components. We usually do not filter this solution. However, if filtration is desired make sure that the membrane filter is stable at high pH. 4. 1 M Ammonium carbonate: dissolve 9.61 g of ammonium carbonate ((NH4)2CO3) in 100 mL of water (w/v). We do not filter this solution. 5. 10 mL plastic tubes. 2.4 Two-Step SolidPhase Extraction

1. dNTP–NTP standard mix: 0.1 μM 2′-deoxycytidine-5′-triphosphate (dCTP), 0.1 μM thymidine-5′-triphosphate (dTTP), 0.1 μM 2′-deoxyadenosine-5′-triphosphate (dATP), 0.1 μM 2′-deoxyguanosine-5′-triphosphate (dGTP), 0.1 μM cytidine-5′-triphosphate (CTP), 0.1 μM uridine-5′-triphosphate (UTP), 0.1 μM adenosine-5′-triphosphate (ATP), and 0.1 μM guanosine-5′-triphosphate (GTP) in sterile water. A volume of 10 μL from the 10 μM stock solution of dNTP standard and 10 μL from the 10 μM stock NTP standard are mixed with 980 μL of sterile water to make the dNTP–NTP standard mix with a final concentration of 0.1 μM each (see Note 4). Store on ice. 2. Isotope-labeled dNTP mix: 10 μM working stock of 13 2′-deoxyadenosine C10,15N5 5′-triphosphate 13 15 13 2′-deoxyguanosine C10,15N5 (dATP C10, N5), 13 15 5′-triphosphate (dGTP C10, N5), 2′-deoxythymidine 13 C10,15N2 5′-triphosphate (dTTP13C10,15N2), and 2′-deoxycytidine 13C9,15N3 5′-triphosphate (dCTP13C9,15N3) prepared in sterile water (see Note 5). 3. Isotope-labeled NTP mix: 100 μM working stock of adenosine 13 C10,15N5 5′-triphosphate (ATP13C10,15N5), uridine 13 C9,15N2 5′-triphosphate (UTP13C9,15N2), guanosine 13C10 15 5′-triphosphate (GTP13C10), and cytidine N3 15 5′-triphosphate (CTP, N3) prepared in sterile water (see Note 5). 4. Muscle sample mix for dNTP and NTP measurement: 12.5 μL of the solution after extraction and neutralization is mixed with 1.25 mL of the eluted solution from the boronate column (see Note 6). Store on ice. 5. Hydrophilic–lipophilic-balanced (HLB) extraction cartridge: Oasis HLB 3 cc, 60 mg (waters). 6. Pre-conditioning solvents: 100% methanol (HPLC grade) and water.

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7. 15 mL conical tubes. 8. Elution solution for HLB: 5% methanol (HPLC grade) in water. 9. Weak anion exchange (WAX) cartridge: Oasis WAX 3 cc, 60 mg, 60 μm, (Waters). 10. Column conditioning solution: 50 mM ammonium acetate buffer, pH 4.5. Dissolve 0.77 g of LC–MS-grade ammonium acetate in 200 mL water and adjust to pH 4.5 with LC–MS– grade acetic acid. Work with acetic acid in a fume hood. Because we use LC–MS-grade reagents, we do not filter this solution. Store at 4 °C in a clean glass bottle. Solution is stable for 4 weeks. 11. Washing solution: 0.5% ammonia aqueous solution in methanol (v/v). To prepare 50 mL of wash solution, add 0.25 mL LC–MS-grade 25% ammonia solution to 49.75 mL HPLC/ LC–MS-grade methanol. Use glass cylinders and tips for measurements. Work with the ammonia solution in a fume hood. Store at 4 °C in a clean glass bottle. Solution is stable for 4 weeks. 12. Disposable culture tubes: borosilicate glass, 12 mm × 75 mm. 13. Elution solution for WAX: methanol/water/ammonia aqueous solution (80/15/5, v/v/v). To prepare 50 mL elution solution, mix 40 mL HPLC/LC–MS-grade methanol, 7.5 mL water, and 2.5 mL LC–MS-grade 25% ammonia solution. Use glass cylinders and tips for measurement. Work with ammonia solution in a fume hood. Store at 4 °C in a clean glass bottle. Solution is stable for 4 weeks. 14. Speed vacuum machine. 15. LC–MS injection vial: 300 μL fixed-insert vials, clear screw top. 16. Sample injection solution: acetonitrile/water/100 mM ammonium acetate, pH 7.7 (30/9/1, v/v/v). To prepare 4 mL sample injection solution, mix 3 mL of LC–MS-grade acetonitrile, 0.9 mL water, and 0.1 mL 100 mM ammonium acetate pH 7.7. Prepare this solution fresh. 17. 100 mM ammonium acetate, pH 7.7: dissolve 1.54 g of LC– MS-grade ammonium acetate in 200 mL water. Adjust to pH 7.7 with LC–MS-grade 25% ammonia solution. Store at 4 °C in a clean glass bottle. We do not filter this solution. Solution is stable for 4 weeks. 2.5 Separation and Detection of dNTPs and NTPs by LC–MS/ MS

1. A SeQuant® ZIC®–cHILIC PEEK column (3 μm, 100 Å, 150 mm × 2.1 mm, Merck).

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2. LC–MS/MS system: Agilent 1290 UHPLC coupled with an Agilent 6490 triple quadrupole mass spectrometer (Agilent Technologies). 3. Mobile phase A: 10 mM ammonium acetate, pH 7.7, in 90/10 water/acetonitrile: to prepare 300 mL of mobile phase A, mix 240 mL water, 30 mL 100 mM ammonium acetate pH 7.7, and 30 mL LC–MS-grade acetonitrile. Because we use LC– MS-grade reagents and solvents, we do not filter the mobile phase. 4. Mobile phase B: 2.5 mM ammonium acetate, pH 7.7, in 90/10 acetonitrile/water: to prepare 400 mL of mobile phase B, mix 360 mL LC–MS-grade acetonitrile, 10 mL 100 mM ammonium acetate pH 7.7, and 30 mL water. 5. Agilent Mass Hunter workstation software.

3

Methods All procedures are carried out at room temperature unless otherwise specified.

3.1 Collection of Muscle Tissue Sample

1. Euthanize animals by cervical dislocation and rapidly dissect out a part (approximately 500 mg) of the thigh muscles (see Note 7). 2. Immediately transfer the dissected tissue into a 1.5 mL Eppendorf tube containing 0.7 mL cold cell lysis solution and freeze in liquid nitrogen (see Note 8). At this stage, the samples can be stored at -80 °C until further analysis.

3.2 Extraction of dNTPs and NTPs from the Muscle Tissue Sample

1. Thaw the frozen muscle tissue sample on ice. Add five stainlesssteel beads to the sample tube and homogenize the sample in the bead beater in the cold room until completely homogenized to a smooth paste (see Note 9). 2. Centrifuge the tubes at 20,000 × g for 5 min at 4 °C to pellet the cell debris (see Note 10). 3. Add 0.8 mL neutralization solution to a new 2 mL Eppendorf tube and add 0.7 mL neutralization solution to another 2 mL Eppendorf tube. Keep the tubes on ice. Transfer the supernatant from step 2 to the first 2 mL tube containing 0.8 mL ice-cold neutralization solution and vortex for 20 s (see Note 11). Separate the layers by centrifugation at 20,000 × g for 1 min at 4 °C. Carefully take 0.7 mL of the upper layer and add it to the second 2 mL tube containing 0.7 mL ice-cold neutralization solution. Vortex for 20 s and separate the layers by centrifugation at 20,000 × g for 1 min at 4 °C. Transfer the upper layer to a fresh 1.5 mL Eppendorf tube and use for both

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the boronate affinity column separation procedure and for preparing the sample mix for solid-phase extraction (see Note 12). 3.3 Separation of dNTPs from NTPs with the Boronate Affinity Column

The following procedures should be carried out under gravity flow. 1. Wash the column with 12 mL of column washing buffer. 2. Condition the column with 12 mL of ambic buffer (see Note 13). 3. Adjust 500 μL of the solution obtained in step 3 under Subheading 3.2 to pH 8–9 using 25 μL of 1 M ammonium carbonate and load onto the boronate column. Discard the flowthrough (see Note 14). 4. Transfer the boronate columns to 10 mL plastic tubes that have been pre-chilled on ice (see Note 15). 5. Elute the dNTPs with 2.5 mL of ambic buffer. 6. Remove columns from collection tubes and wash with 12 mL column washing buffer to regenerate columns (see Note 16).

3.4 Two-Step SolidPhase Extraction

The following procedures are carried out either under gravity flow or by using a vacuum manifold. 1. Add 5 μL of isotope-labeled dNTP internal standard (IS) mix and 5 μL of isotope-labeled NTP internal standard mix to the muscle sample mix and to the dNTP–NTP standard mix prepared in Subheading 2.4. Keep the samples on ice. 2. Pre-condition the HLB cartridges with 2 mL methanol (see Note 17). 3. Pre-condition the cartridges with 2 mL water. 4. Transfer the pre-conditioned cartridges to the 15 mL conical tubes pre-chilled on ice. Load the standard mix and muscle sample mix onto the preconditioned cartridges and collect the flow-through into the tubes. 5. Elute the analytes into the same tubes with 1 mL elution solution for HLB. 6. Adjust the pH of the solution in the conical tubes to pH 4.5 with acetic acid and keep the samples on ice (see Note 18). 7. Pre-condition the WAX cartridges with 2 mL methanol (see Note 17). 8. Pre-condition the WAX cartridges with 2 mL water. 9. Condition the cartridges with 2 mL column conditioning solution. 10. Load the samples obtained in step 6 above onto the conditioned cartridges. Discard the flow-through.

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11. Wash the cartridges with 2 mL wash solution and discard the flow-through. 12. Place the cartridges into a glass tube and elute the analytes with 2 mL elution solution for WAX (see Note 19). 13. Evaporate the solution in a speed vacuum machine for an hour at a temperature below 35 °C. Transfer the remaining solution to an LC–MS injection vial and evaporate to complete dryness (see Note 20). 14. Reconstitute the residue in 50 μL sample injection solution for the HILIC–MS/MS analysis. Samples can be stored at -20 °C for a week until further analysis. 3.5 Separation and Detection of dNTPs and NTPs by LC–MS/ MS

The dNTPs and NTPs are analyzed in the same run using the Agilent LC–MS/MS Mass Hunter Workstation Software. 1. Place the dNTP–NTP standard mix and muscle sample mix vials in the 4 °C auto-sampler. UHPLC separation of analytes is carried out on a SeQuant® ZIC®–cHILIC column in gradient elution mode with a 0.2 mL/min flow rate using mobile phase A and mobile phase B. Apply the stepwise gradient program listed in Table 1. The injection volume is 5 μL. The separation is carried out at 35 °C (see Note 21). 2. The LC–MS/MS instrument should be operated in multiplereaction-monitoring (MRM) mode. The MRM transitions from precursor ions to product ions for all four dNTPs, NTPs, and their corresponding isotope-labeled internal standards are listed in Table 2. 3. Set the in-source parameters for MS as listed in Table 3. 4. After the run, analyze the standard mix and muscle sample mix qualitatively using the Qualitative Analysis of MassHunter Acquisition Data software and quantitatively using the QQQ Quantitative Analysis software. ZIC–cHILIC MRM chromatograms of the four dNTPs and four NTPs in a standard mix

Table 1 HPLC gradient program for nucleotide separation on the SeQuant® ZIC®–cHILIC column Time (min)

Mobile phase A (%)

Mobile phase B (%)

0.0

20

80

7.0

20

80

12.0

40

60

17.0

40

60

19.0

20

80

23.0

20

80

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Table 2 Precursor ions and product ions of the dNTP and NTP analytes used in the analysis

Analyte

Precursor ion [M + H] +

Product ion (m/z)

Collision energy (V)

dCTP

468

112

15

dTTP

483

81

15

dATP

492

136

15

dGTP

508

152

18

CTP

484

112

25

UTP

485

97

25

ATP

508

136

30

GTP

524

152

25

dCTP13C15N

480

119

15

dTTP13C15N

495

86

15

dATP13C15N

507

146

15

dGTP13C15N

523

162

18

15

CTP

487

115

25

UTP13C15N

496

102

25

ATP13C15N

523

146

30

13

534

157

25

GTP

N

C

Table 3 Triple quadrupole mass spectrometer parameters used for the analysis Parameter

Value

QQQ: Gas temperature

200 °C

QQQ: Gas flow

14.1 L/min

QQQ: Nebulizer pressure

20 psi

QQQ: Sheath gas temperature

320 °C

QQQ: Nebulizer gas flow

10.0 L/min

QQQ: Capillary voltage

4000 V

QQQ: Nozzle voltage

400 V

and in muscle sample mix are shown in Fig. 1a, b, respectively. Calculate the concentration of dNTPs and NTPs in muscle sample mix by comparing the isotope-labeled internal standard (IS) response ratio of the analytes in the muscle sample with the

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dATP m/z 492 -> 136

ATP m/z 508 -> 136

dCTP m/z 468 -> 112

CTP m/z 484 -> 112

counts

counts

A

dGTP m/z 508 -> 152

GTP m/z 524 -> 152

UTP m/z 485 -> 97

dTTP m/z 483 -> 81

8

9

10

11

12 13 14 retention time (min)

15

16

17

18

8

9

10

11 12 13 14 retention time (min)

15

16

17

18

15

16

17

18

dATP m/z 492 -> 136

ATP m/z 508 -> 136

dCTP m/z 468 -> 112

CTP m/z 484 -> 112

counts

counts

B

dGTP m/z 508 -> 152

GTP m/z 524 -> 152

UTP m/z 485 -> 97

dTTP m/z 483 -> 81

8

9

10

11

12 13 14 retention time (min)

15

16

17

18

8

9

10

11

12 13 14 retention time (min)

Fig. 1 ZIC–cHILIC–HPLC multiple reaction monitoring (MRM) chromatograms of dNTPs and NTPs in a standard mixture solution (a) and in a muscle tissue extract (b). The MRM transitions for the target analytes are shown

IS response ratio of the analytes in the standard. IS response ratio is calculated as peak area ratio of the analyte to its corresponding internal standard. The result is finally expressed as a ratio of dNTPs to total NTPs in the sample (see Note 22).

4

Notes 1. We use a Bullet Blender Storm homogenizer with air cooling from Next Advance, but any bead beater with a homogenizing option can be used. Work with the homogenizer in a cold room. The stainless-steel beads we use are from Next Advance, but beads from other companies can also be used. 2. Freon has low viscosity and drips very easily during transfer, so care should be taken while preparing the solution. Solution

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preparation should be carried out in a fume hood, and the tips and tubes used with Freon should be collected separately for incineration. Make the solution fresh each time. 3. To pack the column, close the outlet of the column and add 1.0 mL water to mark a line for the following packing. Open the outlet and discard the water. Fill the column with the hydrated gel to the marked line. The packed column can be regenerated and re-used several times before observing any NTP leakage. 4. A 10 mM mixture of the four dNTPs is prepared from 100 mM stock solutions with sterile water and serially diluted in sterile water to make final stock solutions of 10 μM. A mixture of four NTPs is prepared the same way as the dNTP standards. The stock solutions in water can be stored at -20 °C for 6 months. Although these solutions tolerate several freeze–thaw cycles, we divide the stock solutions into small aliquots and use the solutions for no more than three freeze–thaw cycles. 5. A 10 mM mixture of the four isotope-labeled dNTPs is prepared from 100 mM stock solutions with sterile water and serially diluted in sterile water to make final working stock solutions of 10 μM. A mixture of four isotope-labeled NTPs is prepared the same way as the dNTPs to a final working stock solution of 100 μM. 6. The remaining 1.25 mL of eluted solution from the boronate column can be stored at -20 °C for 1 week in case a repeat analysis is required. Because the amount of NTPs is much higher than dNTPs in biological samples, a sample volume of 12.5 μL can provide a strong signal without causing any interference with the dNTP peaks. 7. 300–500 mg of muscle tissue, corresponding to 500 μL volume of an Eppendorf tube, is sufficient for analysis. 8. We have observed that placing the dissected tissue into lysis solution and immediately flash freezing increases the extraction efficiency. 9. For homogenization using stainless-steel beads, be sure to use 1.5 mL safe-lock Eppendorf tubes. Alternately, we use parafilm around the lid of the tubes to ensure tight sealing and to avoid opening of the lid during homogenization. The time for homogenization can vary between 3 and 5 min depending on tissue type and size and the speed settings of the machine. Keep the tubes on ice for 30 s after every 1 min of homogenization to avoid sample overheating. Keep the tubes on ice after homogenization. 10. Remove the parafilm sealing, if used during homogenization, before the centrifugation step. Prepare the neutralization

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solution tubes needed in step 3 beforehand and keep on ice. Work in a fume hood when handling tubes with Freon. 11. Work in a fume hood when handling tubes with Freon. For some tissue types, floating debris builds up after centrifugation. When transferring the supernatant from step 2, care should be taken to avoid collecting any floating debris. During vortexing, make sure the two layers are well mixed giving a milky appearance. 12. Be careful to only remove the upper aqueous phase and avoid taking up any Freon. Store the samples on ice if proceeding immediately. If not, samples can be stored at -20 °C for a week. 13. Add ambic buffer when no more washing buffer drips down. We let the solution flow through under gravity, without applying any extra pressure, to completely remove the washing buffer before loading the ambic buffer. 14. The boronate matrix absorbs the ribonucleotides under alkaline conditions (pH 8.9). No dNTPs or NTPs were found in the flow-through solution when analyzed with HPLC during standardization of the protocol. 15. We make holes in the lids of the tubes to hold the boronate columns. 16. The manufacturer suggests low-pH buffers for eluting the bound NTPs. However, we observed a decreased gel volume when washing with acidic solution so we use washing buffer adjusted to pH 8.9 instead. With this buffer, the NTPs can be quickly removed without shrinking the column. 17. Ensure not to dry the cartridge at any step. A vacuum manifold can be used to regulate the flow when handling many samples. 18. For muscle sample mix, adjust the pH with 50% acetic acid (diluted from 100% LC–MS-grade acetic acid). The volume to set the pH of the sample mix could range between 20 and 30 μL. Use 0.5% acetic acid to adjust the pH of the standard mix. The volume to set the pH of the standard mix is approximately 10 μL. 19. We use cellophane tape to hold the cartridge over the glass tube. 20. Roughly 100–150 μL is left after the first evaporation step. The second evaporation step takes approximately 1 h to completely dry the solution but can be shorter or longer depending on the volume transferred to the LC–MS vial. Ensure complete dryness. 21. In order to minimize contamination of the ion source, each run in the stepwise gradient method should be divided into

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three sections, in which the eluent from the column is diverted to waste from 0 to 7 min and from 19 to 23 min. 22. We have carried out method validation including accuracy, precision, and linearity several times with consistent results; therefore, method validation is not required for every batch of runs. Running quality control samples at the beginning and the end of the sample run to test the sensitivity of the LC–MS/ MS instrument is highly recommended to ensure the accuracy of the analysis.

Acknowledgements This work was supported by the Swedish Cancer Society and the Swedish Research Council. References 1. Reichard P (1988) Interactions between Deoxyribonucleotide and DNA-synthesis. Annu Rev Biochem 57:349 2. Buckland RJ, Watt DL, Chittoor B, Nilsson AK, Kunkel TA, Chabes A (2014) Increased and imbalanced dNTP pools symmetrically promote both leading and lagging strand replication infidelity. PLoS Genet 10(12): e1004846. https://doi.org/10.1371/journal. pgen.1004846 3. Deem A, Keszthelyi A, Blackgrove T, Vayl A, Coffey B, Mathur R, Chabes A, Malkova A (2011) Break-induced replication is highly inaccurate. PLoS Biol 9(2):e1000594. https://doi.org/10.1371/journal.pbio. 1000594 4. Watt DL, Buckland RJ, Lujan SA, Kunkel TA, Chabes A (2016) Genome-wide analysis of the specificity and mechanisms of replication infidelity driven by imbalanced dNTP pools. Nucleic Acids Res 44(4):1669–1680. https:// doi.org/10.1093/nar/gkv1298 5. Gupta A, Sharma S, Reichenbach P, Marjavaara L, Nilsson AK, Lingner J, Chabes A, Rothstein R, Chang M (2013) Telomere length homeostasis responds to changes in intracellular dNTP pools. Genetics 193(4): 1095–1105. https://doi.org/10.1534/genet ics.112.149120 6. Poli J, Tsaponina O, Crabbe L, Keszthelyi A, Pantesco V, Chabes A, Lengronne A, Pasero P (2012) dNTP pools determine fork progression and origin usage under replication stress. EMBO J 31(4):883–894. https://doi.org/10. 1038/emboj.2011.470

7. Schmidt TT, Reyes G, Gries K, Ceylan CU, Sharma S, Meurer M, Knop M, Chabes A, Hombauer H (2017) Alterations in cellular metabolism triggered by URA7 or GLN3 inactivation cause imbalanced dNTP pools and increased mutagenesis. Proc Natl Acad Sci U S A 114(22):E4442–E4451. https://doi.org/ 10.1073/pnas.1618714114 8. Wanrooij PH, Engqvist MKM, Forslund JME, Navarrete C, Nilsson AK, Sedman J, Wanrooij S, Clausen AR, Chabes A (2017) Ribonucleotides incorporated by the yeast mitochondrial DNA polymerase are not repaired. Proc Natl Acad Sci U S A 114(47): 12466–12471. https://doi.org/10.1073/ pnas.1713085114 9. Wanrooij PH, Tran P, Thompson LJ, Carvalho G, Sharma S, Kreisel K, Navarrete C, Feldberg AL, Watt DL, Nilsson AK, Engqvist MKM, Clausen AR, Chabes A (2020) Elimination of rNMPs from mitochondrial DNA has no effect on its stability. Proc Natl Acad Sci U S A 117(25):14306–14313. https://doi.org/10.1073/pnas.1916851117 10. Pontarin G, Gallinaro L, Ferraro P, Reichard P, Bianchi V (2003) Origins of mitochondrial thymidine triphosphate: dynamic relations to cytosolic pools. Proc Natl Acad Sci U S A 100(21):12159–12164. https://doi.org/10. 1073/pnas.1635259100 11. Leanza L, Ferraro P, Reichard P, Bianchi V (2008) Metabolic interrelations within guanine deoxynucleotide pools for mitochondrial and nuclear DNA maintenance. J Biol Chem 283(24):16437–16445. https://doi.org/10. 1074/jbc.M801572200

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12. Sherman PA, Fyfe JA (1989) Enzymatic assay for deoxyribonucleoside triphosphates using synthetic oligonucleotides as template primers. Anal Biochem 180(2):222–226. https://doi. org/10.1016/0003-2697(89)90420-X 13. Ferraro P, Franzolin E, Pontarin G, Reichard P, Bianchi V (2010) Quantitation of cellular deoxynucleoside triphosphates. Nucleic Acids Res 38(6):e85. https://doi.org/10.1093/ nar/gkp1141 14. Wilson PM, Labonte MJ, Russell J, Louie S, Ghobrial AA, Ladner RD (2011) A novel fluorescence-based assay for the rapid detection and quantification of cellular deoxyribonucleoside triphosphates. Nucleic Acids Res 39(17): e112. https://doi.org/10.1093/nar/gkr350 15. Nick McElhinny SA, Watts BE, Kumar D, Watt DL, Lundstrom EB, Burgers PM, Johansson E, Chabes A, Kunkel TA (2010) Abundant ribonucleotide incorporation into DNA by yeast replicative polymerases. Proc Natl Acad Sci U S A 107(11):4949–4954. https://doi.org/10. 1073/pnas.0914857107 16. Williams JS, Lujan SA, Kunkel TA (2016) Processing ribonucleotides incorporated during eukaryotic DNA replication. Nat Rev Mol Cell Biol 17(6):350–363. https://doi.org/10. 1038/nrm.2016.37 17. Nick McElhinny SA, Kumar D, Clark AB, Watt DL, Watts BE, Lundstrom EB, Johansson E, Chabes A, Kunkel TA (2010) Genome instability due to ribonucleotide incorporation into DNA. Nat Chem Biol 6(10):774–781. https://doi.org/10.1038/nchembio.424 18. Di Pierro D, Tavazzi B, Perno CF, Bartolini M, Balestra E, Calio R, Giardina B, Lazzarino G (1995) An ion-pairing high-performance liquid chromatographic method for the direct simultaneous determination of nucleotides, deoxynucleotides, nicotinic coenzymes, oxypurines, nucleosides, and bases in perchloric acid cell extracts. Anal Biochem 231(2): 407–412. https://doi.org/10.1006/abio. 1995.0071 19. Shewach DS (1992) Quantitation of deoxyribonucleoside 5′-triphosphates by a sequential boronate and anion-exchange high-pressure liquid chromatographic procedure. Anal Biochem 206(1):178–182. https://doi.org/10. 1016/s0003-2697(05)80030-2 20. Coquel F, Silva MJ, Techer H, Zadorozhny K, Sharma S, Nieminuszczy J, Mettling C,

Dardillac E, Barthe A, Schmitz AL, Promonet A, Cribier A, Sarrazin A, Niedzwiedz W, Lopez B, Costanzo V, Krejci L, Chabes A, Benkirane M, Lin YL, Pasero P (2018) SAMHD1 acts at stalled replication forks to prevent interferon induction. Nature 557(7703):57–61. https://doi.org/ 10.1038/s41586-018-0050-1 21. Sharma S, Koolmeister C, Tran P, Nilsson AK, Larsson NG, Chabes A (2020) Proofreading deficiency in mitochondrial DNA polymerase does not affect total dNTP pools in mouse embryos. Nat Metab 2(8):673–675. https:// doi.org/10.1038/s42255-020-0264-z 22. Hakansson P, Hofer A, Thelander L (2006) Regulation of mammalian ribonucleotide reduction and dNTP pools after DNA damage and in resting cells. J Biol Chem 281(12): 7834–7841. https://doi.org/10.1074/jbc. M512894200 23. Olafsson S, Whittington D, Murray J, Regnier M, Moussavi-Harami F (2017) Fast and sensitive HPLC-MS/MS method for direct quantification of intracellular deoxyribonucleoside triphosphates from tissue and cells. J Chromatogr B Analyt Technol Biomed Life Sci 1068-1069:90–97. https://doi.org/10. 1016/j.jchromb.2017.10.008 24. Cohen S, Megherbi M, Jordheim LP, Lefebvre I, Perigaud C, Dumontet C, Guitton J (2009) Simultaneous analysis of eight nucleoside triphosphates in cell lines by liquid chromatography coupled with tandem mass spectrometry. J Chromatogr B Analyt Technol Biomed Life Sci 877(30):3831–3840. https:// doi.org/10.1016/j.jchromb.2009.09.030 25. Jia S, Marjavaara L, Buckland R, Sharma S, Chabes A (2015) Determination of deoxyribonucleoside triphosphate concentrations in yeast cells by strong anion-exchange high-performance liquid chromatography coupled with ultraviolet detection. Methods Mol Biol 1300:113–121. https://doi.org/10.1007/ 978-1-4939-2596-4_8 26. Kong Z, Jia S, Chabes AL, Appelblad P, Lundmark R, Moritz T, Chabes A (2018) Simultaneous determination of ribonucleoside and deoxyribonucleoside triphosphates in biological samples by hydrophilic interaction liquid chromatography coupled with tandem mass spectrometry. Nucleic Acids Res 46(11): e66. https://doi.org/10.1093/nar/gky203

Chapter 20 Detection of UV-Induced Deletions in Mitochondrial DNA Gabriele A. Fontana and Hailey L. Gahlon Abstract Mitochondrial DNA (mtDNA) mutations are found in several human pathologies and are associated with aging. Deletion mutations in mtDNA result in the loss of essential genes for mitochondrial function. Over 250 deletion mutations have been reported and the common deletion is the most frequent mtDNA deletion linked to disease. This deletion removes 4977 base pairs of mtDNA. It has previously been shown that exposure to UVA radiation can promote the formation of the common deletion. Furthermore, aberrations in mtDNA replication and repair are associated with formation of the common deletion. However, molecular mechanisms describing the formation of this deletion are poorly characterized. This chapter describes a method to irradiate human skin fibroblasts with physiological doses of UVA and the subsequent detection of the common deletion by quantitative PCR analysis. Key words Mitochondrial DNA, UVA radiation, Deletion Mutations, Common Deletion, Replication and Repair

1

Introduction Several human pathologies and mitochondrial disorders are associated with deletions in mtDNA [1–4]. Deletions can range in size from small (3–100 bp) to large (>1000 bp) [5]. Furthermore, mtDNA is a multi-copy genome present in a state of heteroplasmy, where both wild type and mutated genomes co-exist. If these deleted genomes persist in high copy numbers, they can alter mitochondrial fitness and lower ATP production [6]. While pathways concerning mtDNA replication and repair are linked to the formation of the common deletion [7], fundamental mechanisms to understand the deletion process remain poorly understood. Previous work has shown that UV exposure can induce the formation of mtDNA deletions [8, 9], including the 4977 bp common deletion [10]. Still, important questions remain to elucidate the molecular basis for how UVA can promote mtDNA deletion formation. In this chapter, we describe a method for the irradiation of human skin fibroblasts with UVA. This method relies

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_20, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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on administering physiological doses of UVA that are applied in sequential irradiations to prevent cell death. Furthermore, we provide details for high purity mtDNA isolation, including a key step involving the use of magnetic beads that removes nuclear contamination and significantly increases the purity of mtDNA. Furthermore, to detect and quantify the common deletion, we describe a quantitative PCR approach that is both rapid and reliable and can discriminate the common deletion from wild-type mtDNA.

2

Materials

2.1 UVA Irradiation of Human Skin Fibroblasts

1. Human normal skin fibroblasts cell line BJ-5ta (see Note 1). 2. Culture medium for the BJ-5ta skin fibroblast cell line, with and without supplements (see Note 1). 3. Incubator for mammalian cell culture set at 37 °C with 5% CO2. 4. UVA irradiation device (see Note 2). 5. 15 cm Petri dishes. 6. Soda-lime glass lids for Petri dishes. 7. 1x DPBS (14190-094, Gibco). 8. 15 mL Falcon tubes. 9. Cell scrapers. 10. Centrifuge.

2.2 Isolation of mtDNA

1. QIAprep Spin Miniprep kit (27104, Qiagen). 2. Sonicator: we use a Vibra-Cell Ultrasonic Liquid Processor (Sonics and Material, model VCX750). 3. 1.5 mL Eppendorf tubes. 4. AMPure XP paramagnetic beads (A63880, Beckman Coulter). 5. Vortexer. 6. Magnetic rack for 1.5 mL Eppendorf tubes. 7. 70% EtOH solution. 8. Nuclease-free water. 9. Nanodrop.

2.3 Common Deletion Detection by qPCR

1. GoTaq qPCR master mix (A6001, Promega) see Table 1 for composition of 2x primer mix. 2. Vortexer. 3. Table-top centrifuge. 4. Nuclease-free water. 5. Primers: 100 μM in desalted water (see Table 2 and Note 3).

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6. Ice or metal rack for Eppendorf tubes. 7. Filter tips for pipettes. 8. 0.2 mL and/or 0.1 mL Eppendorf tubes. 9. qPCR device (see Note 4).

3

Methods The method for mtDNA purification presented here is adapted from a previous study [11]. The adaptation to this method aims to increase the final mtDNA yield and to reduce the contamination of nuclear DNA fragments. Overall, these modifications result in an increased accuracy in mtDNA quantification and more robust qPCR data. Dilutions and solutions containing mtDNA and primers should be prepared using DNase-free water. The long-term storage of stock and diluted mtDNA solutions should be at -20 °C and frequent freeze–thaw cycles should be avoided. In addition, primer stock solutions and the qPCR master mix should be stored in frozen aliquots to reduce freeze–thaw exposure. The procedure of mtDNA extraction requires a clean workspace to avoid contamination with nucleases, and while the use of filtered pipette tips is recommended, it is not strictly needed. However, we strongly suggest using filtered pipette tips for the preparation and the assembly of qPCR reactions.

3.1 UVA Irradiation of Human Skin Fibroblasts

1. Passage BJ-5ta human fibroblasts regularly every 2–3 days when reaching a confluency of ~70–80%, and maintain in a DMEM-based culture medium as reported in the American Type Culture Collection (ATCC) guidelines (see Note 1). 2. For each replicate, seed ~107 cells per plate in 15 cm Petri dishes, and be sure to account for the amount of cells needed for both irradiated and non-irradiated plates used as controls. UVA irradiation is usually performed after 2–3 days, when cells reach a confluency of ~80–90%. 3. Prior to irradiation, replace the culture medium with 15 mL DMEM with no additional supplementation (see Note 1). While the presence of phenol red does not affect the UVA irradiation, the absence of FBS is crucial, as some proteins contained in FBS may undergo oxidation and work as photosensitizers. These unwanted chemical reactions can confound the measurements of UVA-dependent biological effects and should, therefore, be avoided. 4. As a control for the conditions of UVA-treated cells, similarly replace the medium of the non-irradiated cells and maintain the non-irradiated cells under the sterile hood during the entire duration of UVA treatment.

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Fig. 1 Scheme of the UVA treatment regime performed on the BJ-5ta human fibroblast cell line. The protocol to induce the formation of the mtDNA common deletion spans 5 days, with an accumulation of 20 J/cm2 UVA per day. Each day, two UVA treatments of 10 J/cm2 UVA were performed in serum- and antibiotic-free medium in an irradiation chamber BS-02 (Opsytec Dr. Grobel, see Note 2) equipped with 8 UVA lamps. With this irradiation chamber and setup, a single dose of 10 J/cm2 UVA is reached in ~25 min. Between the two UVA treatments, a recovery period in complete culture medium is performed. After the last UVA treatment of each day, cells are maintained in complete culture medium to allow recovery overnight. For time-course experiments, cells are harvested by scraping in DPBS from days 3 to 5 (corresponding to accumulated UVA doses of 60–100 J/cm2); for single timepoint experiments, cells are harvested at day 5 (corresponding to an accumulated UVA dose of 100 J/cm2). Cell pellets are washed with DPBS and stored at -20 °C. Following mtDNA extraction, qPCR reactions are performed to monitor the amount of CD-deleted mtDNA within the total mtDNA population

5. Remove the lids of the Petri dishes and cover the dishes with soda-lime glass lids. These lids maintain sterile conditions during irradiation as well as filter out UVB radiation that can be produced from the UVA lamps. This ensures that the cells are only exposed to UVA radiation. If using a dosimeter, cover it with a soda-lime glass lid. 6. In the irradiation chamber BS-02 equipped with 8 UVA-only lamps (see Note 2), set the UVA dose to 10 J/cm2. This UVA dose is reached after ~25 min of irradiation. The UVA treatment regime we follow is depicted in Fig. 1. 7. Retrieve the plates from the UVA device. 8. Remove the medium from the non-treated and UVA-treated cells and replace with 15 mL culture medium. Replace the soda-lime glass lid with plastic lids. 9. Let the cells recover for 2 h in the incubator at 37 °C with 5% CO2. 10. Repeat steps 3–7 to irradiate the cells with another UVA dose of 10 J/cm2. The UVA-treated cells have now accumulated a total UVA dose of 20 J/cm2.

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11. Incubate cells overnight in the incubator. 12. Repeat steps 3–9 the following day until the cells have reached the desired accumulated UVA dose. In this procedure, the cells will accumulate a total of 20 J/cm2 of UVA per day. For the isolation of mtDNA and subsequent detection of the mitochondrial common deletion, we perform either time course experiments, collecting the irradiated cells at days 3, 4, and 5 (corresponding to an accumulated UVA does of 60, 80, and 100 J/cm2, respectively), or a single timepoint at day 5 (corresponding to an accumulated UVA dose of 100 J/cm2). 13. To collect the cells, wash once with 15 mL DPBS and then scrape cells in 8 mL DPBS. After collection in 15 mL falcon tubes, centrifuge at 3000 rpm for 3 min at room temperature and then discard the supernatant. Store pellets at -20 °C until needed (pellets will be stable for several months). 3.2 Isolation of mtDNA

1. Gently thaw the cell pellets on ice. 2. Wash the cells with 1 mL DPBS and transfer the cells suspension to 1.5 mL Eppendorf tubes. Spin at 3000 rpm for 3 min at room temperature and discard the supernatant. 3. Take the AMPure XP paramagnetic beads from the refrigerator and let the suspension equilibrate to room temperature during the first step of mtDNA purification. As the binding of DNA fragments to the beads is temperature-dependent, this equilibration is crucial for reproducibility. 4. Add 250 μL of Buffer P1 from the QIAprep Spin Miniprep kit, with RNAse added. Resuspend the cell pellet by vortexing. 5. To physically lyse the cells, perform sonication. We use a VibraCell Ultrasonic Liquid Processor, with the following protocol: 30 s sonication at maximum intensity, 2 min on ice, repeat five times. 6. Proceed by adding 250 μL of Buffer P2. For subsequent steps, follow the manufacturer’s instructions for the Qiaprep Spin Miniprep kit. In the final step, elute the mtDNA in 50 μL of Buffer EB. 7. Resuspend the AMPure XP paramagnetic beads by vortexing. Add half the volume (from step 6) of the beads (i.e., 25 μL) to the eluted mtDNA solution. Vortex thoroughly to allow binding of mtDNA molecules to the beads. 8. Mount the Eppendorf tubes on a magnetic rack and collect the supernatant. The supernatant contains nuclear genomic DNA fragments and can be saved as a quality control or discarded. 9. Wash the beads twice with 500 μL of 70% EtOH. After the last wash, let the EtOH evaporate by leaving the Eppendorf tubes on the bench at room temperature with the lid open for 5 min.

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Fig. 2 qPCR amplification plots for the genomic ACTB primer pair, with and without beads treatment. A primer pair was designed to hybridize to an intron of the human ACTB gene to indicate the presence of nuclear genomic DNA fragments in mtDNA samples. To decrease nuclear DNA contamination following the mtDNA preparation with the Qiaprep Spin Miniprep kit, an additional purification step with the AMPure XP paramagnetic beads was performed. The graphs show the amplification of the targeted ACTB genomic region from the same cell pellet following mtDNA purification, however, one fraction received additional treatment with beads and one did not. These data represent technical triplicates for both samples (i.e., with and without bead treatment) with 6 ng of mtDNA per reaction as input. The qPCR program encompasses 50 cycles to reveal even “late” amplification products. As shown in the graph, the treatment with beads removes virtually all nuclear genomic contamination from the mtDNA preparation

Avoid excessive drying of the beads as it may result in lower mtDNA recovery. 10. Add 20 μL of nuclease-free water to the beads and vortex thoroughly. Incubate the resuspended beads on the bench at room temperature for 30 min, vortexing every 10 min. This enables detachment of mtDNA molecules from the beads. 11. Mount the Eppendorf tubes on a magnetic rack and transfer the supernatants to new tubes. The supernatant contains the purified mtDNA which, as determined by monitoring a genomic region of the ACTB gene by qPCR (see Fig. 2), is devoid of detectable nuclear genomic DNA contaminants. 12. Measure the mtDNA concentration. We usually measure the mtDNA concentration by nanodrop, with a yield of 50–100 ng/μL mtDNA depending on the initial cell number and treatment conditions. 13. Store the purified mtDNA at -20 °C. This material is stable indefinitely if frequent freeze–thaw cycles are avoided. 3.3 Common Deletion Detection by qPCR

1. Gently thaw the purified mtDNA on ice. Dilute each thawed sample in nuclease-free water to a final concentration of 2 ng/μ L of mtDNA. Briefly spin the tubes containing the diluted mtDNA in a table-top centrifuge and keep on ice. 2. Thaw the primer solutions and the SYBR-green-based GoTaq 2x qPCR master mix on ice. Once thawed, gently vortex and

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briefly spin the tubes in a table-top centrifuge. Keep the tubes on ice. 3. For the following steps, always keep the working samples and reagents on ice or in a pre-cooled metal rack fitted for Eppendorf tubes. To avoid contamination, use filtered pipette tips. 4. Prepare the mtDNA mix. Aliquot 4–6 ng of diluted mtDNA and bring to a final volume of 6 μL with nuclease-free water. As each experimental condition is analyzed for total, CD-deleted and undeleted mtDNA and for genomic ACTB, prepare four aliquots of the same mtDNA mix. In addition, account for technical replicates (we perform 2–3 technical replicates for each sample) and for pipetting errors (we calculate 10–20% additional volume for each reagent). The analysis of genomic ACTB is a quality control for nuclear genomic DNA contamination. Once it is determined that the sample preparation conditions do not yield significant nuclear genomic DNA contamination (see Fig. 2), this control may be omitted. 5. Prepare the 2x primer mix following the instructions reported in Table 1 and using the primers reported in Table 2. The Table 1 Composition of 2x primer mix Total mtDNA GoTaq qPCR master mix

6 μL

Total mtDNA forward primer 100 μM

0.05 μL (0.4 μM final)

Total mtDNA reverse primer 100 μM

0.05 μL (0.4 μM final)

CD-deleted mtDNA

Undeleted mtDNA

6 μL

6 μL

CD-deleted mtDNA forward primer 100 μM

0.05 μL (0.4 μM final)

CD-deleted mtDNA reverse primer 100 μM

0.05 μL (0.4 μM final)

Undeleted mtDNA forward primer 100 μM

0.05 μL (0.4 μM final)

Undeleted mtDNA reverse primer 100 μM

0.05 μL (0.4 μM final)

Genomic ACTB 6 μL

Genomic ACTB forward primer 100 μM

0.05 μL (0.4 μM final)

Genomic ACTB reverse primer 100 μM

0.05 μL (0.4 μM final)

Total volume:

~ 6.1 μL

~ 6.1 μL

~ 6.1 μL

~ 6.1 μL

The indicated volumes are relative to one sample, and the final concentration of primers is calculated based on the total volume of a qPCR sample, i.e., ~12 μL

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Table 2 Primers used in this study. The sequences are provided in the 5′–3′ direction Forward

Reverse

Amplicon

Total mtDNA *

TAGCCCTAAACCTCAACAGT

TGCGCTTACTTTGTAGCC TTCAT

211 bp

CD-deleted mtDNA *

TTCCTCATCACCCAAC TAAAAA

TTCGATGATGTGGTCTTTGG

125 bp

Undeleted mtDNA

CATCTGTACCCACGCCTTCT

TCGATGATGTGGTCTTTGGA

207 bp

CAGCGGAACCGCTCA TTGCCAATGG

295 bp

Genomic ACTB * TCACCCACACTGTGCCCATC TACGA

The primers indicated with * derive from a previous study (Phillips et al. 2017, [12])

localization of primers within the undeleted and CD-deleted mtDNA is reported in Fig. 3 and the melting curves of the primers are depicted in Fig. 4. 6. Add 6 μL of mtDNA mix (from step 4, corresponding to approximately 4–6 ng mtDNA) to each tube containing the four different 2x primer mixes. Gently vortex and spin samples in a table-top centrifuge. 7. The qPCR program is performed as follows in a Rotor-Gene 6000 qPCR device (see Note 4): • Polymerase activation: 95 °C for 2 min, 1 cycle. • Amplification and acquisition: denaturation at 95 °C for 20 s, annealing and extension at 60 °C for 40–60 s, 40–50 cycles. At the end of this step, acquire SYBR green fluorescence. • (Optional) Dissociation (or melting) curve: ramp temperature from 60 to 99 °C, with an increase of 1 °C per min. Acquire SYBR green fluorescence each minute (see Note 5). 8. Export the Ct values corresponding to each sample. We usually use a threshold of 0.2 normalized fluorescence to obtain Ct values. Calculate the mean values among the technical replicates from the same biological sample. 9. For each biological sample, calculate the ΔCt for the CD-deleted and undeleted mtDNA species following these formulas: • ΔCtCD-deleted = mean CtCD-deleted – mean Cttotal mtDNA • ΔCtundeleted = mean Ctundeleted – mean Cttotal mtDNA 10. For each corresponding UVA-treated and untreated sample, calculate the ΔΔCt for the CD-deleted and undeleted mtDNA species following these formulas:

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Fig. 3 Localization of the mtDNA CD and of the qPCR primer pairs used to quantify relative CD levels in the mtDNA pool. (a) Schematic representation of human mtDNA, a 16.6 kb circular molecule encompassing genes encoding key factors of the electron transport chain (ETC) complexes (colored boxes), rRNAs (yellow boxes), tRNAs (black boxes, tRNAs names not indicated for clarity), two origins of replication (black arrows) and non-coding regions (white boxes). The two strands of the mtDNA, named heavy (H, black line) and light (L, grey line) strands, are indicated. The CD affects a region of 4977 bp and ablates six genes encoding ETC proteins and five tRNAs. The CD is flanked by two 13 bp-long direct repeats (purple and green ovals) named CD5´ and CD3´. (b) Localization of the three primer pairs used in qPCR reactions to quantify the relative abundance of CD-deleted molecules (right panel) among the undeleted mtDNA pool (left panel). The total mtDNA primer pair (green arrows) is localized in the 12 rRNAs gene of both mtDNA species. Conversely, the primer pair used to monitor undeleted mtDNA molecules (orange arrows) localizes in the ND5 gene and flanks the CD3´ direct repeat. The primer pair that reveals the presence of CD-deleted mtDNA molecules (blue arrows) flanks the CD3´ direct repeat and amplifies the distinctive deletion junction region that originates from the splicing of parts of the ND5 and ATP6/8 genes

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Fig. 4 Melting curves of the qPCR primer pairs. (a) Melting curves of the primer pairs used to quantify the total mtDNA, undeleted and CD-deleted mtDNA molecules. The curves represent technical triplicates for each primer pair and data were derived from the same mtDNA sample purified with paramagnetic beads. Each sample contains 6 ng of mtDNA per reaction and qPCR was performed after 40 cycles prior to the melting analysis. (b) Melting curves of the primer pairs used to quantify the nuclear genomic ACTB contamination. The same qPCR conditions described in panel a were followed, with the exception that this analysis of the mtDNA preparation was not subjected to beads treatment

• ΔΔCtCD-deleted UVA-treated = ΔCtCD-deleted – mean Ctuntreated • ΔΔCtundeleted UVA-treated = ΔCtundeleted – mean Ctuntreated 11. For each UVA-treated sample, calculate the fold change (FC) of CD-deleted and undeleted mtDNA species following these formulas: • FCCD-deleted UVA-treated = 2^ - (ΔΔCtCD-deleted UVA-treated) • FC undeleted UVA-treated = 2^ - (ΔΔCtundeleted UVA-treated) 12. Calculate the relative percentage of CD-deleted and undeleted mtDNA species in each UVA-treated sample using the following formulas: • Sum FC = FCCD-deleted UVA-treated + FCundeleted UVA-treated • % CD-deleted in UVA-treated = (FCCD-deleted UVA-treated/ SUM FC) × 100% undeleted in UVA-treated = (FC undeleted UVA-treated / SUM FC) × 100

4 Notes 1. BJ-5ta cells (CRL-4001) were obtained from the ATCC. These cells are human fibroblasts isolated from neonatal foreskin and immortalized by expression of hTERT. These cells grow adherently and are maintained under Biosafety Level 1 regulations. BJ-5ta cells are cultured in a medium composed of four parts of Dulbecco’s Modified Eagle’s Medium (DMEM 1x + GlutaMAX, 31966-021, Gibco), one part of Medium 199 (22340020, Gibco), 10% fetal bovine serum (FBS, SH30066.03HI, GE Healthcare), and 0.01 mg/mL Hygromycin B (SH30066.03HI, Sigma). Hygromycin B is a selection

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antibiotic needed to maintain the expression of hTERT. For UVA treatments, the BJ-5ta cells are maintained in DMEM with no supplements for the duration of the irradiation. 2. For UVA irradiation, we use the irradiation chamber BS-02 (Opsytec Dr. Grobel), equipped with 8 UVA lamps, a UV– MAT device to program the irradiation regime and a dosimeter monitoring the accumulated UVA dose in real-time. This device allows for controlled UVA irradiation that can be performed in a dose- and time-controlled manner. The dosecontrolled program allows one to set a specific UVA dose that is monitored in real-time by the internal dosimeter. Irradiation stops once the programmed dose is reached. As reported in the technical specifications of the device, the UVA lamps may produce a small proportion of UVB. In order to avoid the unwanted UVB radiation exposure on the cells, we place soda-lime lids over the Petri dishes containing the cells for the duration of the irradiation. This step ensures both sterile working conditions as well as the ability of the soda-lime glass to filter out the unwanted UVB radiation, but still allowing the passage of UVA. To match these conditions and to measure only UVA, we also cover the dosimeter with a soda-lime glass lid. 3. Primers were purchased from Eurogentec at a concentration of 100 μM in desalted water, aliquoted, and stored at -20 °C. The primer sequences and relative amplicons are reported in Table 2. Primer sequences for total mtDNA, CD-deleted mtDNA and genomic ACTB were used as described previously [12]. The primers for undeleted mtDNA were designed specifically for this study. The localization of primers in the undeleted and CD-deleted mtDNA species is depicted in Fig. 3 and the melting curves obtained are presented in Fig. 4. 4. For qPCR, we use a Rotor-Gene 6000 (Corbett Research) device, equipped with 2 gears that allow for the performance of qPCR using 36 or 72 samples in 0.2 mL or 0.1 mL tubes, respectively. The data are collected, visualized, and exported to a Microsoft Excel spreadsheet using the Rotor-Gene 6000 Series Software 1.7 (Corbett Research). 5. The dissociation curve calculates the melting temperature of the qPCR-generated amplicons, which in this setup is calculated as the temperature at which the SYBR green fluorescence abruptly decreases as a consequence of the dissociation of the dye from the double-stranded PCR products. This analysis is generally used to reveal primer specificity, controlling for off-target annealing, as only one melting point value is expected for each qPCR reaction (see, for example, Fig. 4).

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Acknowledgements This work was supported by Federation of Migros Cooperatives; ETH Zurich Federation in association with the World Food System Center at ETH Zurich. References 1. Dimauro S, Davidzon G (2005) Mitochondrial DNA and disease. Ann Med 37(3):222–232. h t t p s : // d o i . o r g / 1 0 . 1 0 8 0 / 07853890510007368 2. Russell O, Turnbull D (2014) Mitochondrial DNA disease-molecular insights and potential routes to a cure. Exp Cell Res 325(1):38–43. https://doi.org/10.1016/j.yexcr.2014. 03.012 3. Taylor RW, Turnbull DM (2005) Mitochondrial DNA mutations in human disease. Nat Rev Genet 6(5):389–402. https://doi.org/ 10.1038/nrg1606 4. Yuan Y, Ju YS, Kim Y, Li J, Wang Y, Yoon CJ, Yang Y, Martincorena I, Creighton CJ, Weinstein JN, Xu Y, Han L, Kim HL, Nakagawa H, Park K, Campbell PJ, Liang H, Consortium P (2020) Author Correction: Comprehensive molecular characterization of mitochondrial genomes in human cancers. Nat Genet. https://doi.org/10.1038/s41588-0200629-y 5. Lott MT, Leipzig JN, Derbeneva O, Xie HM, Chalkia D, Sarmady M, Procaccio V, Wallace DC (2013) mtDNA variation and analysis using Mitomap and Mitomaster. Curr Protoc Bioinformatics 44:1 23 21-26. https://doi. org/10.1002/0471250953.bi0123s44 6. Lu J, Sharma LK, Bai Y (2009) Implications of mitochondrial DNA mutations and mitochondrial dysfunction in tumorigenesis. Cell Res 19(7):802–815. https://doi.org/10.1038/cr. 2009.69 7. Fontana GA, Gahlon HL (2020) Mechanisms of replication and repair in mitochondrial DNA deletion formation. Nucleic Acids Res 48(20):

11244–11258. https://doi.org/10.1093/ nar/gkaa804 8. Powers JM, Murphy G, Ralph N, O’Gorman SM, Murphy JE (2016) Mitochondrial DNA deletion percentage in sun exposed and non sun exposed skin. J Photochem Photobiol B 165:277–282. https://doi.org/10.1016/j. jphotobiol.2016.10.030 9. Torregrosa-Munumer R, Goffart S, Haikonen JA, Pohjoismaki JL (2015) Low doses of ultraviolet radiation and oxidative damage induce dramatic accumulation of mitochondrial DNA replication intermediates, fork regression, and replication initiation shift. Mol Biol Cell 26(23):4197–4208. https://doi.org/10. 1091/mbc.E15-06-0390 10. Berneburg M, Grether-Beck S, Kurten V, Ruzicka T, Briviba K, Sies H, Krutmann J (1999) Singlet oxygen mediates the UVA-induced generation of the photoagingassociated mitochondrial common deletion. J Biol Chem 274(22):15345–15349. https:// doi.org/10.1074/jbc.274.22.15345 11. Quispe-Tintaya W, White RR, Popov VN, Vijg J, Maslov AY (2013) Fast mitochondrial DNA isolation from mammalian cells for nextgeneration sequencing. BioTechniques 55(3): 1 3 3 – 1 3 6 . h t t p s : // d o i . o r g / 1 0 . 2 1 4 4 / 000114077 12. Phillips AF, Millet AR, Tigano M, Dubois SM, Crimmins H, Babin L, Charpentier M, Piganeau M, Brunet E, Sfeir A (2017) Singlemolecule analysis of mtDNA replication uncovers the basis of the common deletion. Mol Cell 65(3):527–538 e526. https://doi. org/10.1016/j.molcel.2016.12.014

Chapter 21 Determination of the Ribonucleotide Content of mtDNA Using Alkaline Gels Choco Michael Gorospe, Bruno Marc¸al Repoleˆs, and Paulina H. Wanrooij Abstract Impaired mitochondrial DNA (mtDNA) maintenance, due to, e.g., defects in the replication machinery or an insufficient dNTP supply, underlies a number of mitochondrial disorders. The normal process of mtDNA replication leads to the incorporation of multiple single ribonucleotides (rNMPs) per mtDNA molecule. Given that embedded rNMPs alter the stability and properties of the DNA, they may have consequences for mtDNA maintenance and thereby for mitochondrial disease. They also serve as a readout of the intramitochondrial NTP/dNTP ratios. In this chapter, we describe a method for the determination of mtDNA rNMP content using alkaline gel electrophoresis and Southern blotting. This procedure is suited for the analysis of mtDNA in total genomic DNA preparations as well as in purified form. Moreover, it can be performed using equipment found in most biomedical laboratories, allows the simultaneous analysis of 10–20 samples depending on the gel system employed, and can be modified for the analysis of other mtDNA modifications. Key words Ribonucleotides, rNMPs, Alkaline gels, Denaturing gels, Alkaline hydrolysis, Southern blot

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Introduction Mature mitochondrial DNA (mtDNA) was shown to contain incorporated ribonucleotides (rNMPs) as early as the 1970s [1– 3]. The far majority of these ribonucleotides are present as single rNMPs inserted during mtDNA replication or repair [4–6]. The rNMP content of mtDNA is partly determined by the size and balance of the cellular dNTP pool, as indicated by altered rNMP frequency and identity when cellular dNTP levels are perturbed in yeast cells or in mammalian tissues [6, 7]. Moreover, mtDNA rNMP content and identity were found to be altered in patientderived fibroblasts and mouse models carrying disease-associated defects in enzymes involved in mitochondrial nucleotide

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_21, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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metabolism [5, 8]. The rNMP content of mtDNA can therefore serve as an indication of intramitochondrial NTP/dNTP ratios, and can be used for this purpose either together with a method for determining cellular dNTP and NTP pools (such as the one described in Chapter 19 of this volume), or alone, e.g., when the size or nature of the sample precludes direct measurement of nucleotide pools. Finally, even single incorporated rNMPs can change the local structure and elasticity of DNA and increase the risk of strand breaks by several orders of magnitude [9–12]. The frequency of rNMPs in mtDNA may thereby have direct implications for many DNA-related processes, such as transcription, replication, and repair [13], and is interesting to study in its own right. For these reasons, it can be of interest to experimentally determine the rNMP content of mtDNA. Several next-generation sequencing-based technologies have been developed for mapping the location and identity of rNMPs genome-wide, thus also including the mtDNA [14–17]. While these methods do not per se provide quantitative information of mtDNA rNMPs, they can be modified to do so [18]. However, the sequencing-based methods rely on a chain of optimized enzymatic steps, e.g., for DNA amplification, ligation of adapters, and other modifications prior to sample sequencing, as well as an established bioinformatic pipeline and expertise for data analysis. Here, we describe an alternative procedure for the determination of mtDNA rNMP content using alkaline gels and Southern blotting. This method is robust, relatively simple to perform using equipment available in most biomedical laboratories, and allows the parallel analysis of 10–20 samples in a single experiment (depending on the gel system available). Like many of the sequencing-based approaches, the described procedure exploits the reactive nature of the hydroxylgroup on carbon-2 of the ribose moiety of rNMPs that sensitizes the adjacent phosphodiester bond to hydrolysis under alkaline conditions. Alternatively, the protocol can be used in conjunction with enzymatic cleavage at embedded rNMPs by RNase H2, yielding comparable results [7]. The alkali-treated DNA is electrophoretically separated under denaturing conditions when DNA migration is only determined by the length, not the shape, of the single-stranded DNA fragments. The DNA is transferred to a porous membrane and the mtDNA visualized using a 32P-labeled probe. In a quantification step adapted from the previous work [19], the migration of the alkali-treated DNA is compared to that of the untreated DNA sample and a DNA ladder run in parallel, allowing the approximate determination of the number of rNMPs per single strand of mtDNA. If single-stranded DNA probes are employed in the Southern blot, this method allows the strandspecific analysis of mtDNA rNMP content. This procedure can be modified to determine the frequency of any mtDNA modification, provided a robust treatment exists for

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the specific nicking of mtDNA at the site of the modification of interest. Depending on the type of starting material, the amount of DNA required for this procedure varies from ca 600 ng of pure or enriched mtDNA to up to 10 μg of total genomic DNA from mammalian tissues with low mtDNA copy number.

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Materials Prepare all reagents using ultrapure water (prepared by purifying deionized water to attain sensitivity of 18.2 MΩ-cm at 25 °C) and analytical grade reagents. Prepare and store all reagents at room temperature unless indicated otherwise. Handle all reagents, especially the strong bases and acids, with the appropriate care and according to the manufacturer’s instructions.

2.1 Alkaline Hydrolysis

1. 5 M Sodium hydroxide (NaOH). 2. 0.5 M ethylenediaminetetraacetic acid (EDTA), pH 8.0: weigh 186.1 g of EDTA and transfer it to a 1 L beaker or bigger. Add 800 mL water to the beaker and stir. Adjust the pH to 8.0 with 5 M NaOH (see Note 1). Make up to 1 L with water and transfer to a 1 L screw-cap bottle or bigger. Autoclave. 3. 1 M Tris–HCl, pH 8.0: Autoclave. 4. Tris–EDTA (TE) buffer: 0.5 M EDTA, pH 8.0, 1 M Tris–HCl, pH 8.0. Autoclave. 5. 5 M Potassium hydroxide (KOH). 6. 5 M Acetic acid. 7. 6x Alkaline loading buffer: 300 mM NaOH, 6 mM EDTA, 18% (w/v) Ficoll 400, 0.15% (w/v) bromocresol green, 0.25% (w/v) xylene cyanol. To make 5 mL, combine 300 μL 5 M NaOH, 60 μL 0.5 M EDTA, 0.9 g Ficoll, 7.5 mg bromocresol green, and 12.5 mg xylene cyanol. Make up to 5 mL with water.

2.2 Alkaline Gel Electrophoresis

1. Neutral agarose solution: 2% in water (see Note 2). Just before use, weigh 2 g agarose and transfer to a 250 mL glass bottle or Erlenmeyer flask, add 100 mL water. Microwave to boiling, swirling occasionally, and ensure all agarose has melted. 2. Alkaline agarose solution: 0.7% agarose, 1 mM EDTA, 30 mM NaOH. About 30–60 min before planned use, weigh 3.5 g of agarose and transfer to a 1 L glass bottle or Erlenmeyer flask. Add 500 mL water and weigh, noting the weight of the bottle/ flask together with its contents. Microwave until boiling, then adjust to original weight with water. Allow to cool on a magnetic stirrer until flask is cool enough to hold (see Note 3). Add

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3 mL of 5 M NaOH and 1 mL 0.5 M EDTA, and stir gently to mix before pouring the gel. 3. Alkaline run buffer: 30 mM NaOH, 1 mM EDTA. A few hours prior to use, mix 15 mL of 5 M NaOH and 5 mL of 0.5 M EDTA in 2.5 L of water. Place in cold room to cool. 4. DNA marker mix: 0.33 ng/μL 1 kb DNA ladder, 8.33 mM EDTA and 1x alkaline loading buffer in TE (see Note 4). Prepare an aliquot sufficient for loading two lanes just prior to use by mixing 2 μL of a 5 ng/μL dilution of 1 kb DNA ladder, 1 μL of 0.5 M EDTA, and 6 μL 6x alkaline loading buffer in 21 μL of TE buffer. 5. Agarose gel electrophoresis system with a 20 cm × 20 cm gel tray and 30-well comb 1.5 mm thick. 2.3

Gel Treatments

1. Clear glass dish large enough to accommodate the gel (e.g., glass lasagna dish). 2. 2 M Tris–HCl, pH 7.2: weigh 242.3 g of Tris base in a 1 or 2 L beaker; add water to 800 mL. Stir until dissolved. Using fuming HCl, bring the pH to 7.2. Note that a large volume of HCl is required (>50 mL). Make up to 1 L with water (see Note 5). 3. Neutralization solution: 1 M Tris–HCl, pH 7.2, 2 M NaCl. Weigh 233.76 g of NaCl and transfer to a 2 L bottle. Add 1 L of 2 M Tris–HCl, pH 7.2 and adjust the volume to 2 L with water. 4. 50x TAE: 50 mM EDTA, 2 M Tris, 1 M glacial acetic acid. Weigh 242 g of Tris-base and dissolve in 700 mL of water. Add carefully 57.1 mL of 100% glacial acetic acid and 100 mL of 0.5 M EDTA, pH 8.0. Bring the final volume to 1 L with water. 5. 1x TAE with 0.5 μg/mL ethidium bromide (EtBr): add 10 mL of 50x TAE to 490 mL water and transfer to a bottle covered with aluminum foil. Add EtBr to a final concentration of 0.5 μg/mL (see Note 6).

2.4

DNA Transfer

1. Hybond-N+ positively charged nylon transfer membrane. 2. 3 MM Whatman paper. 3. 20x SSC: 3 M NaCl, 0.3 M sodium citrate dihydrate. Dissolve 350.8 g of NaCl and 176.4 g of sodium citrate in 1.5 L water. Bring the total volume to 2 L with water. 4. 10x SSC: combine 500 mL of 20x SSC and 500 mL of water; mix and store in a glass bottle. 5. 3x SSC: combine 150 mL of 20x SSC and 850 mL of water; mix and store in a glass bottle.

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6. Platform (e.g., large Eppendorf tube rack, or a glass or plexiglass plate). 7. CL-1000 Ultraviolet (UV) crosslinker. 2.5 Southern Hybridization

1. Prehybridization solution: 1 mM EDTA, 7% SDS in 250 mM phosphate buffer, pH 7.2. Dissolve 30.25 g of sodium phosphate monobasic (NaH2PO4·H2O) in 500 mL water to make 0.5 M sodium phosphate monobasic solution. Dissolve 35.95 g of disodium phosphate (Na2HPO4·2H2O) in 500 mL water to make 0.5 M sodium phosphate dibasic solution. Take 400 mL of the sodium phosphate dibasic solution and bring the pH to 7.2 with the sodium phosphate monobasic solution (approximately 100–150 mL). To 500 mL of the resulting 500 mM phosphate buffer at pH 7.2, add 70 g of SDS and 2 mL of 0.5 M EDTA. Heating the solution will aid in dissolving the SDS. Adjust the final volume to 1 L with water. 2. Radioactively labeled DNA probe and DNA ladder (see Notes 7 and 8). 3. Radioactively labeled DNA ladder (see Notes 8 and 9). 4. Hybridization oven. 5. Wash buffer 1: 3x SSC, 0.1% SDS. 6. Wash buffer 2: 0.3x SSC, 0.1% SDS. 7. BAS–MS imaging plate (Fujifilm) and appropriate cassette. 8. Typhoon 9400 imager (GE Amersham) or equivalent.

2.6

Quantification

1. Image J software (NIH). 2. Spreadsheet application, such as Microsoft Excel.

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Methods All procedures, unless specified, are carried out at room temperature.

3.1 Alkaline Hydrolysis

1. For each sample to be analyzed, transfer 3–5 μg of total genomic DNA (3 μg for DNA from muscle tissue or cultured cells, 5 μg for liver or heart) or 300–500 ng of pure mtDNA to each of two 1.5 mL Eppendorf tubes (see Notes 10–11). These will constitute the untreated and alkali-treated samples. Add 0.5 μL of 0.5 M EDTA to the DNA samples (see Note 12) and adjust the total volume to 23.5 μL with TE buffer. Mix the solution well by tapping the tube or vortexing at low speed (shaking setting #4–5) (see Note 13). Spin down the sample briefly.

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2. Add 1.5 μL of 5 M KOH to each of the samples to be alkalitreated (but not to untreated controls). Vortex immediately at low speed to evenly distribute the KOH, and spin down briefly. 3. Incubate both the control and the alkali-containing samples in a pre-warmed incubator at 55 °C for 2 h (see Note 14). 4. Stop the alkaline hydrolysis by adding 2.2 μL of 5 M acetic acid to the alkali-treated (but not to control) samples. Vortex at low speed to distribute acetic acid evenly. 5. Add 6 μL of 6x alkaline loading buffer to all samples. Mix by flicking or vortexing at low speed; spin down briefly. 3.2 Alkaline Gel Electrophoresis

Preparation of the gel (steps 1–2) and of the alkaline running buffer (under Subheading 2.2, item 3 in the Materials) is ideally done during the 2-h alkaline hydrolysis reaction. 1. Cast a neutral base layer for the gel by pouring 50–100 mL of boiled 2% agarose in water into a gel tray (20 cm × 20 cm). Leave to solidify. 2. When the base layer has set, and the alkaline gel solution cooled down enough to allow addition of the NaOH and EDTA to complete the mixture, insert the comb into the gel tray and pour the alkaline gel. Set aside to solidify. 3. Place the set gel in the gel tank in the cold room, and fill with cold alkaline run buffer until just covered. Carefully remove the comb (see Note 15). 4. Load 15 μL of DNA ladder mix in the outermost wells on each side of the gel (see Notes 16 and 17). Then, load the entire volume of all the untreated and then all the alkali-treated samples in the remaining wells. Do not leave frequent empty wells between samples (see Note 18). 5. Run at 46 V for 16–17 h (see Note 19).

3.3

Gel Treatments

1. After the run, transfer the gel to a large glass dish. If required to fit the gel in the dish, cut off a few centimeters from the bottom of the gel. 2. Neutralize the gel by incubating it with neutralization solution for 40 min while shaking gently at 40 rpm (see Note 20). 3. Aspirate the neutralization solution, rinse the gel once with distilled water, and stain by incubating the gel in 1x TAE with 0.5 μg/mL EtBr for 30–60 min. 4. Transfer the gel to a tray or clean dish, and collect the 1x TAE with EtBr for reuse (see Note 21). Image the gel using a UV-light source and camera or Chemidoc® Touch imaging system (see Note 22). With the help of a ruler and scalpel, cut the gel ca 0.5 cm above the RNA species that are typically

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Fig. 1 Setup of DNA transfer from gel to nylon transfer membrane. See the text for details

visible in the untreated samples, and discard the lower part of the gel. 3.4

DNA Transfer

In this step, the DNA will be transferred from the gel to a nylon membrane using the setup depicted in Fig. 1. 1. Cut out 3 MM Whatman paper (three pcs.) and Hybond-N+ positively charged nylon transfer membrane (one pc) that exceed the dimensions of the gel by 0.5 cm along each axis. In addition, cut out two long strips of 3 MM Whatman paper that measure 0.5 cm wider than the gel and are long enough to reach the bottom of the glass dish when placed on top of the platform (see Fig. 1). 2. Pour 10x SSC into the glass dish to a height of ca 3 cm. Across the top of the glass dish, place a solid platform that is larger than the gel, and at least as long but narrower than the dish. This will leave gaps on both sides of the platform so the long Whatman papers can access the 10x SSC in the dish below. We use a large Eppendorf tube rack that fits perfectly on top of the glass dish (see Note 23). In addition, pour 10x SSC into another dish or tray. 3. Overlay the two long Whatman papers on each other, briefly wet them in the extra dish of 10x SSC, and place them on top of the platform (see Note 24). Carefully roll out bubbles, e.g., using a 25 mL pipette (see Note 25). 4. Overlay two of the shorter Whatman papers, briefly wet them in 10x SSC, and place them centrally on the platform, on top of the longer strips of Whatman paper. Roll out bubbles.

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5. Invert the agarose gel so it is facing downward in your hands, and place it centrally on top of the shorter pieces of Whatman paper (see Note 26). Peel off the thin layer of high-percentage neutral agarose (see Note 27). Roll out bubbles. 6. Remove one of the paper covers of the transfer membrane, label the top right corner of the membrane with soft pencil, remove the other paper cover, and briefly dip the membrane in 10x SSC (see Note 28). Place the transfer membrane exactly on top of the alkaline gel. Once it has touched the gel, the position of the membrane should not be adjusted. Carefully roll out bubbles. 7. Briefly wet the last short piece of Whatman paper, and place it on top of the membrane. Make sure the transfer membrane is fully covered; roll out bubbles. 8. Use two strips of parafilm or cling film to seal off both long edges of the dish (Fig. 1). The strips should not overlap the smaller piece of Whatman placed uppermost in the stack; rather, their purpose is to prevent the paper towels that will form the next layer from coming in direct contact with the long strips of Whatman or the 10x SSC. 9. Place a thick pile (15–20 cm) of paper towels on top of the short Whatman paper (see Note 29). Place another platform or glass plate on top of the stack; add a weight (e.g., 250 mL flask of buffer) to keep the layers stable and compact. Incubate overnight, or up to 48 h. 10. After transfer, remove the paper towels and invert the smaller, central stack (2xWhatman/gel/membrane/Whatman) on a clean surface. Peel off the two small pieces of Whatman paper to expose the gel that is still on top of the membrane. Use a soft pencil to mark the position of some of the wells on the membrane, then peel the gel off of the membrane. Place the membrane in a dish with 3x SSC (see Note 30). 11. Place the transfer membrane in a UV cross-linker and crosslink at a dose setting of 200,000 μJ/cm2 (see Note 31). Place the transfer membrane back into the 3x SSC. It can be stored in the buffer for up to a few weeks, or in dry form (wrapped in cling film) practically indefinitely. If stored dry, rehydrate the membrane briefly in 3x SSC before proceeding to the next step. 3.5 Southern Hybridization

Handling of radioactive materials should only be performed by trained personnel. Follow the rules and regulations in place at your institute. 1. Roll up the transfer membrane with the DNA facing inward, and place in a hybridization tube. Add 25 mL of pre-warmed

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prehybridization solution and incubate under rotation in a hybridization oven at 65 °C for 30–120 min (see Note 32). 2. Denature the labeled probe and DNA ladder at 95 °C for 5 min; cool on ice for 5–10 min. Centrifuge briefly. 3. Transfer the used prehybridization solution to a 50 mL Falcon tube and add 25 μL of the labeled DNA probe and 2.5 μL of the labeled DNA marker (or as found suitable for your probe of choice). Mix by inverting the tube repeatedly, then pour into the hybridization tube with the membrane in it. Incubate under rotation in the hybridization oven at 65 °C for between 4 h and overnight. 4. Collect and store the hybridization solution (see Note 33). Briefly wash the membrane with 25 mL of pre-warmed wash buffer 1. Collect the used wash buffer (see Note 34). 5. Wash the nylon transfer membrane by adding 25 mL of pre-warmed wash buffer 1 and incubating under rotation in the 65 °C hybridization oven for 20 min (see Note 35). 6. Wash the nylon transfer membrane with 25 mL of pre-warmed wash buffer 2, incubating under rotation in the 65 °C hybridization oven for 20 min. Repeat. 7. Wrap the membrane in two layers of cling film (see Note 36). Expose the membrane to a BAS–MS imaging plate in a suitably sized cassette at room temperature overnight or as required (see Note 37). 8. Scan the imaging plate in a Typhoon 9400 imager or equivalent, and save as a .gel file (Fig. 2) (see Note 38). 3.6

Quantification

The data analysis is performed using ImageJ software and Microsoft Excel. The commands and short-cuts described are for the Macintosh versions. The equivalent of the “Command” key in a Windows operating system is “Ctrl”. Should the commands not work in your operating system, use the Help section of the software or internet searches to identify the corresponding commands. 1. Open the .gel image file in ImageJ software. Use the “Rotate” command (Image ! Transform ! Rotate) to rotate the gel, so that the wells are exactly horizontal (see Note 39). With the rectangular selection tool selected (which is default), click-drag to frame a rectangular area of the blot encompassing all the samples, and reaching vertically from just below the wells to as far down as possible on the blot without reaching off the blot (see Note 40). The selected area should preferably include some empty lanes for quantification of background signal. Crop the image (Image ! Crop). Save a copy for future reference (File ! Save as ! jpg).

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Fig. 2 Scanned image of an example Southern blot hybridized with a dsDNA probe for a region of the Cox1 gene in mtDNA. Genomic DNA samples isolated for a previous study [7] from the heart, tibialis anterior (TA) muscle, and the liver of three 16-week-old wild-type mice (1–3, and) were either left untreated (left half of blot) or were treated with alkali (right half of the blot). Two lanes containing DNA ladder (MW1 and MW2) flank the samples; the size of the DNA ladder bands is indicated in kbp

2. Click-drag to frame a rectangular selection (known as a region of interest, or ROI) reaching from top to bottom of the leftmost lane (Fig. 3a). Adjust the width of the ROI to exclude signal from the neighboring lane. Click-drag in the middle of the ROI to overlay it on the most narrow and skewed lanes on the blot; decrease the width further if necessary in order to exclude signal from neighboring lanes. Once a suitable size for the ROI is found, care should be taken not to unintentionally adjust its dimensions when moving it between lanes, as the same size of ROI must be used throughout the analysis. 3. Move the ROI back to the left-most lane. Press OptionCommand-K (Ctrl-Alt-K for Windows) to plot the intensity of the signal over the entire length of the lane. Copy the quantification data by clicking More ! Copy 1st data set (alternatively: Data ! Copy 1st data set), and paste into cell A2 of a new Excel worksheet (Fig. 3b). This analysis essentially divides the signal in the lane into numerous small segments of a defined size. The first column of data in the spreadsheet reflects the migration of DNA in each segment (in cm, measured from the top of the ROI; referred to as “migration” from now on) and the second column contains the signal intensity values in each lane segment. Label the second column (in cell B1) with

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Fig. 3 Quantification of signal intensity using ImageJ software, see the text for details. (a) Rectangular selection tool is used to frame a region of interest (ROI) that reaches from the top to the bottom of lane 1. The width of the ROI is adjusted to exclude signal from neighboring lanes. (b) Signal intensity along the length of the ROI is plotted by pressing “Option-Command-K”. In the plot window, the source data is copied by clicking on “More ! Copy 1st data set”. The procedure exemplified here for lane 2 is repeated for each individual lane, including the DNA ladder lanes and at least one empty lane. (c) View of Excel spreadsheet containing the uppermost rows of the raw signal from each lane (columns B-V) alongside the migration (column A). Background-corrected signal for the first segment (row 2) of sample 1 (MW1) is calculated by subtracting the raw signal of the first segment (row 2) of the empty lane (Bg; cell V2) from the first segment of the raw signal of sample 1 (cell B2) using the formula displayed in cell Y2. (d) The sum of background-corrected signal for sample 1 (column Y) is computed below the last row of data (here on row 1180) using the formula “=sum (Y2:Y1179)”. The formula is then copied to adjacent columns

the name of the sample. Return to ImageJ, close the plot window, and use the right arrow to move the ROI to the next lane (see Note 41). Repeat the quantification for each lane, including the ones with DNA ladder and a minimum of one

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empty lane that will be used to determine the level of background signal. In the spreadsheet, delete all columns corresponding to migration data except for column A. 4. If your version of the spreadsheet software uses commas (“,”) instead of points (“.”) as decimal separators, select all data (Command-A; Ctrl-A in Windows) and replace all decimal points (“.”) with commas (“,”) using Find ! Replace ! Replace all. Furthermore, under Subheading 3.6, step 10, replace the decimal points in the formulas with decimal commas. 5. If several empty lanes were quantified, calculate the average background signal to be used for background correction: label a new column as “background”. In the cell on row 2 of that column, calculate the average of the row-2 signals of all quantified empty lanes. Fill the formula down to all subsequent rows by dragging the lower right corner of the cell downward until the last row of data is reached. 6. To calculate background-subtracted signal intensities: in an empty cell on row 2, enter “=B2-$[column of background signal]2” (Fig. 3c). In the provided example, the background signal is in column V, so the formula used is “=B2-$V2” (see Note 42). Click-drag on the lower right-hand corner of the cell to fill the formula to neighboring cells on the right (to complete the calculation for row 2 of all samples), then fill down until the last row of data (see Note 43). Below the last row of background-subtracted data, calculate the sum of signal intensities in each respective column (Fig. 3d; in our example, the quantified signal data reaches until row 1179 and the sum is computed in row 1180). 7. To calculate normalized signal intensity (% of signal in each segment relative to total signal in the lane): in row 2 of a new column, divide the background-subtracted signal in row 2 of sample 1 by the sum of signal for sample 1, and multiply by 100 (see Note 44). In our example, where the backgroundsubtracted signal for sample 1 is in column Y, and the sum of the signal in row 1180, the formula is: “=Y2/Y$1180*100”. Fill the same formula to cells corresponding to all samples and rows, as done above. 8. To define the “median” migration of each sample: in row 2 of a new column, calculate the cumulative sum of the normalized signal of sample 1 from row 2 onward (see Note 45). In our example, where the normalized signal of sample 1 is in column AU, we use: “=sum(AU$2:AU2)”. Fill the formula to adjacent cells on the right and below, as done above. Scroll down, column by column, and make a note of the row number, where the cumulative sum first reaches the value 50 for each respective sample. This row number, when used together with

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the migration in column A, reflects the median migration of the DNA fragments in each sample. For example, for a sample whose cumulative sum of signal reaches 50 in row 500, the “median migration” value is returned by typing “=A500”. 9. To generate a standard curve using the two DNA ladder lanes (referred to as “MW1” and “MW2”): copy the migration (column A) and the normalized signal intensity data of MW1 and MW2 (from step 7) to three adjacent empty columns (using Paste Special ! Values). Insert a scatter plot with the normalized signal intensity of MW1 and MW2 on the y-axis, and migration on the x-axis (Fig. 4a). Using the x-coordinates of the MW1 peaks in the plot as an initial guide, locate the exact row corresponding to the highest signal within each marker band (i.e., the migration of the “tip” of the peak; see Note 46). Note the corresponding migration value in a table, next to the known size of that band of the DNA ladder (Fig. 4b). Repeat for each individual band in MW1 and MW2. Calculate the log10 of the size of the ladder bands (in nt). Create a scatter plot with these log10 values on the y-axis and the migration values of MW1 and MW2, respectively, on the x-axis (Fig. 4c). Left-click on a point of the MW1 standard curve, choose “Add trendline” and check the boxes for “Display equation on chart” and “Display R-squared value on chart”. Similarly, add a trendline and display the equation for the MW2 standard curve. Because the migration distance of a DNA fragment (“x” in the standard equation) is inversely proportional to the logarithm of its length (“y”), these equations can be used in the next step to convert the median migration of each sample to median length. 10. To calculate the median length of fragments in each lane: use the equations of the standard curves for MW1 and MW2 (from step 9) to convert the median migration values determined for each sample in step 8 into median length. To reduce error introduced by small differences in migration between the two edges of the gel during electrophoresis, the samples on the left half of the gel are compared to MW1, and those on the right half to MW2. Thus, for each individual sample, use the respective standard curve equation, substituting the median migration (defined in step 8) for “x”, to compute the median length by entering: “=10^([equation, with the median migration substituted for x])”. For example, for the MW1 equation (y = -0.1553x + 4.215) and the Untreated heart sample 1 with a median migration corresponding to row 500, the median length is retrieved by entering: “=10^(0.1553*A500 + 4.215)” (Fig. 4d). In contrast, the corresponding alkali-treated sample—with a median migration corresponding to row 692—was run in the right half of the gel, and will thus employ the MW2 equation (y = -

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Fig. 4 Generating a standard curve, see the text for details. (a) A scatter plot of normalized signal intensity for the two DNA ladder lanes MW1 (in blue) and MW2 (in orange) on the y-axis and migration (from column A or CM) on the x-axis. Hover over the peaks to see the approximate migration reading for each peak. (b) The exact

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0.1661x + 4.2489). The median length of the alkali-treated heart 1 sample is calculated by entering: “=10^(0.1661*A692 + 4.2489)”. 11. To calculate rNMP frequency and rNMP content: comparison of the median lengths of untreated and alkali-treated sample allows the determination of rNMP frequency according to the following formula [19]: “=a/((a/b) - 1)”, where “a” and “b” are the median sizes of the untreated and alkali-treated sample, respectively (Fig. 4e). The rNMP frequency reflects the median distance between two rNMPs in the analyzed mtDNA, and can be used to determine the mtDNA rNMP content (the number of rNMPs per single strand of mtDNA) by dividing the genome length by the rNMP frequency (e.g., 16 299 nt in mouse) (Fig. 5).

Fig. 5 rNMP content (rNMPs per single strand of mtDNA) of mtDNA in the heart, tibialis anterior (TA) muscle and liver of 16-week-old wild-type mice. (a) mtDNA rNMP content in the indicated tissues of three individual mice (data from mouse 1, 2, and 3 are shown in blue, orange and grey, respectively). (b) Median mtDNA rNMP content of the data in panel a). Error bars represent standard error; N = 3

ä Fig. 4 (continued) migration of each band corresponding to MW1 (column DC) and MW2 (not shown) is noted next to the known band size in nucleotides (column DB). Log10 of the band size is calculated in the adjacent column (column DD). (c) Standard curve with the log10 of the band size on the y-axis and migration on the xaxis for MW1 (in blue) and MW2 (in orange). Right-click on a data point to add a trendline, then display the equation and R2 value on the plot (in blue and orange for MW1 and MW2, respectively). (d) The standard curve equations for MW1 and MW2 are used to convert the median migration of untreated (in blue) and alkali (KOH) -treated (in orange) samples to median length. For the untreated heart sample 1 (cell DL3; highlighted) with a median migration defined by the value in cell A500, the median length is calculated according to the MW1 equation, with “A500” substituted for “x”. (e) Ribonucleotide (rNMP) frequency for heart sample 1 is computed in cell DL10 (green frame) using the median lengths of the untreated sample (cell DL3; blue frame) and the alkali (KOH) -treated sample (cell DO3; red frame) according to the displayed formula

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Notes 1. EDTA does not readily dissolve in water even after prolonged stirring. It is important to adjust the pH of the solution to 8.0 with 5 M NaOH to increase the solubility. 2. While not essential, we use this 2% agarose solution in water to cast a supportive base for the alkaline gel. This will greatly facilitate handling of the gel after electrophoresis and prevents gel breakage. 3. For good quality gels, it is important that all the agarose has melted. Furthermore, the agarose will disintegrate if heated in an alkaline solution. It is, therefore, important to allow the agarose solution to cool until the bottle is cool enough to touch before adding the NaOH. Stir gently for a few minutes after addition of NaOH and EDTA to ensure complete mixing, but avoid formation of bubbles. Even with these precautions, certain brands of agarose are not compatible with alkaline gels—if the alkaline gel turns yellowish and appears incompletely set after pouring, try another brand of agarose. We use research-grade agarose for DNA electrophoresis from SERVA. 4. Choose a DNA ladder with a suitable range of band sizes, preferably reaching from 250 bp to 10 or 20 kbp. We prepare small aliquot of a 5 ng/μL dilution of the ladder in TE, and store in the freezer for this purpose. This circumvents the need to thaw the stock tube each time to withdraw a very small volume. 5. If carrying out this protocol repeatedly, it can be practical to prepare 2 L of the 2 M Tris–HCl pH 7.2 at a time so as to have an extra 1 L ready for when more neutralization solution needs to be made. 6. Ethidium bromide is a potential carcinogen. Use appropriate care when handling it, and dispose of waste according to local regulations. It can potentially be exchanged for less toxic DNA stains, such as GelRed. 7. Double-stranded DNA (dsDNA) probes ca 400–600 bp in length can be prepared by PCR-amplifying the desired region from gDNA. The oligonucleotide sequences we use for amplifying DNA probes targeting the mtDNA and the nuclear DNA are listed in Table 1. For preparation of the probe, we set up four 50-μL PCR reactions per probe using a standard Taq polymerase (according to manufacturer’s instructions), amplify for 40 cycles with an annealing temperature of 57 °C, and gel-purify the PCR product. Alternatively, a single-stranded oligonucleotide can be used as a probe. This has the advantage of permitting strand-specific determination of rNMP content,

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Table 1 Oligonucleotides used for amplification of dsDNA probes targeting mtDNA (Cox1) or nuclear DNA (18S rDNA region) in mouse (Mus musculus) Oligonucleotide

Sequence

mmCox1 5554F

5′-gag gct ttg gaa act gac ttg tcc c-3′

mmCox1 6009R

5′-ggt ccc ctc ctc cag cgg-3′

mm18S F

5′-gac tca aca cgg gaa acc tca ccc g-3′

mm18S R

5′-cag cgc tcc gcc agg gcc-3′

but often results in a lower signal-to-noise ratio than dsDNA probes. 8. Long dsDNA probes and the DNA ladder can be labeled with α-dCTP32 using a kit containing random primers, Klenowfragment and the appropriate buffer. We use Prime-It II Random primer labeling kit (Agilent technologies, cat #300385 according to manufacturer’s instructions), and remove unincorporated α-dCTP32 using Illustra™ Microspin™ G-25 columns (GE healthcare). Short oligonucleotides are conveniently end-labeled using ɣ-ATP32 and polynucleotide kinase. Non-radioactive procedures for Southern blotting also exist, e.g., ones based on digoxygenin-labeled probes, and have the additional advantage that probes can be stored and used over long periods of time without loss of signal. In our experience, however, the current 32P-based protocol results in cleaner and crisper images and is therefore our method of choice. 9. The DNA ladder used here should be the same one as was used for gel electrophoresis. 10. The amount of DNA to be used per reaction can be modified depending on the starting material and the desired signal strength. We find that 3 μg of total genomic DNA (gDNA) from mouse skeletal muscle (based on DNA concentration measurements using Nanodrop) gives optimal signal, and that ca 5 μg of gDNA from liver or heart is required to obtain a similar signal intensity because of the somewhat lower mtDNA copy number in these tissues. When pure or enriched mtDNA is used, the DNA amount used can be decreased to 300–500 ng. 11. We routinely use gDNA isolated from solid mammalian tissues by incubation with proteinase K followed by phenol–chloroform extraction [7], although grinding of frozen samples could also be considered. Regardless of the isolation protocol, RNase A treatment should be avoided, as it may result in cleavage at incorporated rNMPs under certain conditions. For this reason,

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our DNA preparations still contain RNA species, which we cut off the gel as described in Note 22. 12. Prior to exposure to alkaline conditions, EDTA should be added to the DNA to a total concentration of at least 10 mM to avoid the precipitation of magnesium hydroxide that retards the movement of the DNA in the gel. 13. Vortexing the gDNA at high speed causes the DNA to shear. Therefore, take care to mix gDNA-containing mixtures with care either by flicking the tube or vortexing on a low setting, and use wide-bore or cut tips for pipetting whenever feasible. We cut off the end of 200 μL tips at the 10-μL mark for pipetting gDNA-containing solutions. 14. Use an incubator or a PCR machine with a heated lid. The use of a heating block is not recommended, as condensation of water on the lid will increase the KOH concentration in the sample mixture to above 30 mM. If the incubator is equipped with a shaking function, use low speeds ( T-specific mtZFN monomer (MTM25) recognizes the mutation site and dimerizes with mtZFN (WTM1). Dimerization leads to DNA cleavage, followed by degradation of the mutant mtDNA

the nucleus [3]. Evidence for this directionality of DNA transfer towards the nucleus is seen today in the form of long stretches of nuclear DNA with near perfect homology to mtDNA, commonly referred to as nuclear mitochondrial DNA [nuclear-encoded mitochondrial sequences (NUMTs)], which can confound accurate genotyping of mtDNA by introducing a nuclear bias [4] (Fig. 1a). The first part of this protocol describes a systematic approach to avoid potentially confounding NUMTs while measuring mtDNA heteroplasmy by a judicious selection of the mtDNA genotyping assay used for experiments. While the localization of mtDNA in the mitochondrial matrix may serve as a distinct advantage for the in situ synthesis of OXPHOS subunits, it comes at the cost of increased mutagenesis when compared to nuclear DNA [5]. This increased rate of mutagenesis, as compared with the nucleus, is thought to contribute to the de novo emergence of a plurality of mtDNA species harboring aberrations which can be acquired in somatic tissues during the organism’s life or be inherited if the maternal germline was affected [6]. When discussing this multitude of mtDNA species, the concept of heteroplasmy (coexistence of more than one genotype) has

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been introduced, which is typically expressed as a percentage, as opposed to the familiar concept of alleles used for nuclear genetics. Aberrations of mtDNA can come in the form of single-nucleotide polymorphisms or deletions, and many are known to be pathogenic; however, a disease phenotype is usually only observed at heteroplasmies over 60–90%, with this observed threshold varying depending on the nature of the mutation and the affected tissues [3]. Indeed, most individuals typically harbor basal levels of mtDNA heteroplasmy, at rates that are far below these pathogenic thresholds [7]. It is fortuitous that some repair mechanisms are absent for mtDNA, in particular, effective DNA double-strand break (DSB) repair, which proves beneficial as it opens the door to a viable therapeutic strategy. According to this strategy, the introduction of a mutant-specific DSB in mtDNA results in rapid degradation of mtDNA molecules harboring a pathogenic mutation, and upon the recovery of the initial mtDNA copy number, the heteroplasmy of a cell is shifted to below the pathogenic threshold. Remodeling the mitochondrial genotype by targeted degradation thus leads to a rescue of the phenotype as the heteroplasmic proportion of pathogenic mtDNA decreases (Fig. 1b). Selective degradation of mtDNA can be achieved by several types of designer nucleases, which could specifically be engineered to preferentially anneal to and linearize only mtDNA harboring pathogenic mutations [8]. The easily programmable CRISPR/ Cas9 system does unfortunately not lend itself toward mtDNA targeting as a robust mechanism of RNA delivery into mitochondria has not been documented in mammals [9], thus prohibiting the co-delivery of the guide RNA to the mitochondrion. It is, however, completely feasible to utilize the aforementioned mitochondrial protein import machinery to deliver protein-based DNA modifying factors, such as zinc finger nucleases (ZFNs), transcription activator-like effector nucleases, meganucleases or restriction endonucleases. Our laboratory focuses on engineered mitochondrially targeted ZFNs (mtZFNs), which have been successfully used to degrade mutation-bearing mitochondrial DNA both in vivo [10] and in vitro [11–13], resulting in a shift in the genetic makeup of affected mitochondria and subsequently to the phenotypic rescue. These chimeric nucleases are designed as a pair of constructs and bind to their DNA target site in a tail-to-tail orientation. Each construct encodes a modified nucleolytic domain of the FokI restriction enzyme on the C-terminus linked to an array of Cys2His2 zinc finger DNA binding motifs, with each motif capable of recognizing three base pairs on a DNA double helix. The nucleolytic domains of the FokI enzyme have been modified to act as obligate heterodimers, thus reducing toxicity and enhancing specificity [14, 15]. Multiple Cys2His2 zinc finger domains are strung together into arrays to achieve target site specificity, preventing unspecific degradation of wild-type mtDNA molecules

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with well-optimized arrays being able to discriminate DNA sequences based on a single base pair. Mitochondrial delivery of these constructs is achieved by the N-terminal addition of a mitochondrial transport sequence (MTS) and nuclear export signals (NES), the latter being required to offset the endogenous affinity for zinc finger domains toward nuclear DNA [16]. The second part of the protocol presents the necessary steps to verify and qualify mtZFNs to be used in vivo on mouse models harboring single-nucleotide mtDNA mutations. This chapter follows a similar general workflow to a chapter published previously by our laboratory for in vitro manipulation of human cell lines [17]. It is assumed that the user is in possession of a mouse model harboring mtDNA mutations and that a mouse embryonic fibroblast (MEF) cell line has been isolated and immortalized from this mouse. You will also require a library of zinc finger arrays to screen using this procedure, which can be obtained using several published methods. A simple and commonly used approach is to proceed by modular assembly, which involves selecting zinc finger motifs from previously established and characterized libraries. The zinc finger modules in these libraries bind to pre-defined DNA triplets, which allows them to be assembled into arrays corresponding to longer target sequences [18, 19]. While zinc finger motifs largely behave in a modular fashion, residues in adjacent motifs are known to interact with the target sites of their neighbors in a phenomenon referred to as target-site overlap. To account for this, more laborious approaches exist, which allow for the generation of active and specific arrays with a higher certainty of positive outcome, notably context-dependent assembly [20] and oligomerized pool engineering [21]. Useful background information on zinc finger motifs and their use as DNA recognizing domains can be found here [22, 23]. This protocol will be demonstrated using mtZFNs optimized in an immortalized MEF cell line isolated from the m.5024C > T mt-tRNAAla mouse [24] (Fig. 1).

2

Materials

2.1 Genotyping Design by Pyrosequencing

1. PyroMark® Assay Design software

2.2 Cloning ZF Arrays into mtZFN Expression Vectors

1. KOD Hot Start DNA Polymerase kit. 2. pTracer/CMV/Bsd (modified not to contain BamHI in cycle3GFP) with an mtZFN monomer. 3. pcmCherry with a mtZFN monomer (Addgene: 62803). 4. EcoRI-HF.

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5. BamHI-HF. 6. T4 DNA Ligase. 7. Agarose. 8. SYBR Safe gene staining solution. 9. DH5α subcloning competent cells. 10. Ampicillin. 11. SOC medium. 12. LB Broth. 13. LB Agar plates with 50 μg/mL ampicillin. 14. PCR Clean-up kit. 15. Gel extraction kit. 16. Mini prep kit. 17. Midi prep kit. 2.3 MEF Transformation and Sorting

1. MEF2 kit from Lonza. 2. Cell strainer (50 μL). 3. 5 mL Polystyrene FACS tubes. 4. BD Influx™ Cell Sorter or equivalent.

2.4

DNA Analysis

1. DNA extraction kit from mammalian cells. 2. PyroMark® advanced reagents. 3. PyroMark® Q48 Magnetic Beads. 4. PyroMark® Absorber Strips. 5. PyroMark® Q48 machine. 6. PyroMark® software. 7. KOD Hot Start DNA Polymerase kit. 8. Primers (Table 1). 9. SYBR Green 2X Master Mix. 10. 96-well Optical Real time PCR plates. 11. 96-well Real-time thermal cycler.

Table 1 Primers for heteroplasmy measurement by pyrosequencing Primer name

Primer sequence 5′ ! 3′

PCR F

ATATACTAGTCCGCGAGCCTTA

PCR R

[Biotin] – GCAAATTCGAAGGTGTAGAGAAA

Sequencing primer

AAGTTTAACTTCTGATAAGG

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Table 2 Primers for copy number measurement by qPCR Primer name

Primer sequence 5′ ! 3′

HK2 M. musculus F

GCCAGCCTCTCCTGATTTTAGTGT

HK2 M. musculus R

GGGAACACAAAAGACCTCTTCTGG

ND1 M. musculus R

CCGGCTGCGTATTCTACGTT

ND1 M. musculus F

CTAGCAGAAACAAACCGGGC

12. 96-well Real time Optical seals. 13. qPCR primers (Table 2). 2.5 Common Laboratory Equipment

1. PCR Tubes. 2. 0.5 mL centrifuge tubes. 3. 1.5 mL centrifuge tubes. 4. Standard PCR thermal cycler. 5. DNA mini gel tank. 6. Power Pack. 7. Spectrophotometer. 8. Tube roller. 9. Tilt platform. 10. Benchtop centrifuge. 11. UV transilluminator.

2.6 Non-commercial Reagents

3

1. MEFs with the m.5024C > T mutation.

Methods

3.1 Designing Primers for the Measurement of Heteroplasmy

Due to the high throughput nature of mtZFN heteroplasmy shifting optimization, we recommend creating a pyrosequencing assay which allows for cost effective, rapid, and quantitative genotyping of mtDNA heteroplasmy. In this section, we will present the design of primers for such a pyrosequencing assay that avoids bias from homologous NUMTs as well as observed biases in the pyrosequencing assay itself. 1. Open the mtDNA sequence for the background of the mouse model you are using, in this case C57BL6/N. Identify the position of the heteroplasmic polymorphism and copy 1000 bp both up- and downstream of this position into a text file.

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2. Paste the portion of the mtDNA sequence from step 1 into the Pyromark assay design software. Select your variable base as well as the two bases directly adjacent to it in the pasted sequence as your region of interest and run the primer design (see Note 1). 3. The software will produce a list of trios of primers: two are for creating an amplicon as a template for the pyrosequencing reaction (one will be biotinylated and the other not) and the third is a sequencing primer which will anneal near the position of interest. The assays will be ranked by quality score. Retain the five best primer sets in terms of score (see Note 2). 4. To check for possible nuclear off targets, paste the mtDNA region amplified by each of your prospective amplification primers into the online BLAST NCBI tool [25] and align the amplicon with the murine representative genome (Select the following options: Refseq Representative Genomes—Mus musculus—somewhat similar sequences). 5. The algorithm should return a perfect homology with the complete murine mitochondrial genome as well as any highly homologous nuclear sequences. If any nuclear sequences have perfect or near-perfect homology to the binding site of the sequencing primer, then the primer set should be discarded (see Note 3). 3.2 Cloning Zinc Finger Arrays into Expression Vectors

We routinely use two plasmids encoding mtZFNs with cytoplasmic fluorescent reporters to identify transformed cells. The zinc finger array sites are conveniently flanked by single cutter restriction sites (EcoRI and BamHI), allowing for the rapid substitution of arrays into compatible constructs. The remainder of the constructs includes the FokI obligate heterodimer domain, an MTS, an NES, and an epitope tag as mentioned in the introduction. This section describes the cloning procedures required to substitute the zinc finger arrays. 1. If the DNA sequences encoding your Zinc Finger arrays are not already flanked by an EcoRI restriction site on the 5′ end and a BamHI site on the 3′ end, amplify the DNA with primers containing these restriction sites making sure to preserve the correct orientation and reading frame of the array. Use a 50 μL KOD Hot Start Polymerase PCR reaction by following the manufacturer’s guidelines. 2. Add loading dye and run the PCR reaction in a 1% agarose gel stained with SYBR Safe at 7 V/cm for 1 h along with a 1 kb Plus DNA ladder. 3. View the gel on a UV transilluminator and verify that specific Zinc Finger array bands were produced at the expected DNA length. Excise the correct bands with a clean sharp scalpel and

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recover the amplified DNA from the gel fragment with a commercial Gel extraction kit, eluting the DNA in 43 μL of molecular grade water. 4. Set up restriction reactions in separate PCR tubes for both of your backbone vectors as well as your amplified Zinc Finger arrays. On ice, combine 1 μg of the mtZFN-pTracer and mtZFN-pcmCherry with 20 U of EcoRI-HF and BamHI-HF (1 μL each) and 5 μL of CutSmart® buffer and add molecular grade water up to 50 μL. For the PCR-amplified zinc finger arrays from the previous step, similarly, add 5 μL of CutSmart buffer and the two restriction enzymes to achieve a total volume of 50 μL. Quickly spin down and incubate the samples at 37 °C for 1 h. 5. Add loading dye to digestion products and run in a 1% agarose gel stained with SYBR Safe at 7 V/cm for 1 h along with a 1 kb Plus DNA ladder. 6. View the gel on a UV transilluminator and excise the correct DNA fragments from the gel using a clean sharp scalpel. Recover the amplified DNA from the gel fragment with a commercial Gel extraction kit. Elute the DNA in water and measure the concentrations by spectrophotometry. 7. Use T4 DNA ligase to ligate the backbones with the corresponding Zinc Finger arrays following the manufacturer’s guidelines (see Note 4). 8. Thaw DH5α competent cells on ice and transform them with 5 μL of the ligation reaction from the previous step according to the manufacturer’s guidelines. 9. After transformation by thermal shock at 42 °C, add the total reaction volume to 1 mL of SOC media in a 15 mL conical tube and incubate at 37 °C in an incubator with a shaking platform set to approximately 120 rpm for 1 h. 10. Spread 300 μL of each of the SOC cultures on LB Agar plates supplemented with 50 μg/ml of ampicillin with a sterile spreader and incubate overnight at 37 °C. 11. The next day, pick several clones with a sterile pipette tip from each of your culture plates and grow the individual clones at 37 °C with agitation overnight in 5 ml of LB broth supplemented with 50 μg/ml of ampicillin (see Note 5). 12. Perform a plasmid mini prep from the 5 mL cultures of plasmid according to the manufacturer’s instructions and verify by Sanger sequencing using a CMV Forward primer to ensure that the plasmids contain the correct Zinc Finger array insert in the correct reading frame. 13. Grow successful clones overnight in 100 mL of LB Broth supplemented with 50 μg/ml of ampicillin.

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14. Isolate the plasmid DNA using an endotoxin-free midi prep kit according to the manufacturer’s instructions and resuspend in a small volume of molecular grade water, typically 1 μg/μL as measured by spectrophotometry. Verify the plasmids by sequencing once more prior to beginning experiments (see Note 6). 3.3 MEF Transformation by Electroporation and FACS Enrichment

To ensure that cells have been transformed with both mtZFNencoding plasmids, double positive fluorescence cells must be enriched by FACS (Fig. 2). While several transformation options can be used, one of the most efficient methods of transient transformation for MEF cells is by electroporation. In this protocol, we describe the use of the Lonza Nucleofector II apparatus for MEF transformation, for which we have optimized a procedure which is different than that recommended by the manufacturer’s protocol. Cell lines harboring mtDNA mutations can often require

Fig. 2 Fluorescence gating strategies for this example as measured by a BD Influx™ Cell Sorter. Each point on the scatter plot represents a single event registered by the cytometer. (a) P1 denotes the selection of MEF cells using Side and Forward scatter profiles. (b) P2 gate used to select monodisperse cells and exclude doublets to ensure only positive cells are collected. (c) Untreated MEF cells selected with gates P1 and P2 plotted for GFP and mCherry-specific fluorescence. (d) Cells transformed only with WTM1-pTracer plasmid hence expressing Cycle3GFP. Events in the ‘pTracer’ gate indicated on the scatter plot were selected. (e) Cells transformed only with MTM25-pcmCherry. Events in the ‘mCherry’ gate indicated on the scatter plot were selected. (f) Cells co-transformed with WTM1-pTracer and MTM25-pcmCherry. Events in the ‘Double positive’ gate indicated on the scatter plot were selected

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supplementation with 50 μg/ml of uridine; however, most MEF cell lines can be readily grown at standard conditions (37 °C, 5% CO2) using DMEM supplemented with 10% FCS (v/v), 4.5 g/L d-glucose, Glutamax and 100 mg/L sodium pyruvate. The necessity for uridine can be discussed with the provider of the MEF cell line. When you first receive and expand your MEF cell lines, genotype them by pyrosequencing to establish the heteroplasmy value and freeze down several vials prior to keeping them in continuous culture. Mitochondrial heteroplasmy can drift over time, in which case a vial of cells with the original heteroplasmy can be revived. 1. One day before electroporation, seed cells into a 175 cm2 flask at a density of 1.2 × 107 cells and leave to attach (see Note 7). 2. Prepare plasmid DNA mixtures with 5 μg of each of your conjugated mtZFN containing plasmids in advance in PCR tubes (see Note 8). In this example, the single fluorescence controls are singly transfected mtZFN plasmids (Fig. 2d, e); however, empty vector plasmids can also be used as a control. 3. The following day, detach cells with Trypsin and separate 2 × 106 cells into 15 mL conical tubes. Spin tubes at 90 g for 10 min. Preserve cells for continuous culture and for an untransfected control. 4. Add 2 mL warm media to 6-well plate for each transformation condition, keeping one well for the untreated control. 5. Transform each condition using protocol T-020 on the Nucleofector II apparatus following the manufacturer’s recommendations, but using the total volume of the DNA mixture prepared in advance. 6. The following day, aspirate media and wash cells with 1 × PBS. Aspirate PBS and detach cells with 100 μL Trypsin for 5 min. 7. Prepare a 5 mL round-bottom polypropylene tube for each condition by setting the caps of the tubes aside and placing a CellTrics 50 μm strainer in them. Pass several mL of 1 × PBS through the strainers to avoid cell loss. Discard the PBS from the tube. 8. Add 400 μL of DMEM to each of the wells in step 6 to inhibit the Trypsin, then ensure that as many cells are detached and separated as possible by gently pipetting up and down across over the entire surface of the 6-well plate. 9. Transfer the total volume of each well into a polypropylene FACS tube by passing the cells through the pre-soaked strainer. Label the tube with the corresponding condition and secure the cap. 10. Begin by loading the untreated control cells into your cytometer. Use these cells to establish the gating for your

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subsequent samples. The first two FACS gates should exclude debris and cellular aggregates using the side scatter and forward scatter profiles (Fig. 2a, b). 11. Set fluorescence gating for GFP (excited with a 488 nm laser with 530/30 bandpass and 502 long-pass filters) and mCherry (excited with a 561 nm laser and detected by PMT 610/20 bandpass and 600 long-pass filters). Adjust the PMT values for the fluorescence gating to have the auto-fluorescent cell population near the origin of the scatter plot (Fig. 2c). 12. Collect single and double positive cells from each of your experimental and control conditions into sterile 1.5 mL Eppendorf tubes containing 500 μL of DMEM +10% FBS (Fig. 2d–f). If this is your first attempt at transfecting the MEF cells using your constructs, sort your positive cells two separate tubes, allowing one sample to be analyzed by Western blotting (see Note 9). 13. Centrifuge recovered cells in table-top centrifuge for 10 min at 100 g and aspirate most of the medium in a sterile laminar flow hood, leaving ~50 μL to ensure the cell pellet is not disturbed. 14. Resuspend each cell pellet in 100 μL of warm DMEM and plate in individual wells of a 96-well plate (see Note 10). Once cells have reached confluency, passage the sorted cells into a larger well. 3.4 Analysis of MEF Cells and mtZFN Qualification

The final section describes how to rapidly assess the activity of mtZFNs in MEF cell mitochondria by genotyping. Once expression and import of the constructs is verified, DNA from treated MEFs can be analyzed by pyrosequencing and qPCR to observe heteroplasmy shifts and mtDNA copy number recovery, respectively (Fig. 3).

3.4.1 Measuring Shifts in mtDNA Heteroplasmy

1. Collect cells from the second well of the FACS enriched culture and spin them at 300 g for 5 min. 2. Extract DNA using a commercial kit following the manufacturer’s specifications, eluting the DNA in molecular grade water. Quantify DNA by spectrophotometry. 3. In triplicate for each sample, set up 25 μL PCR reactions to amplify mtDNA for Pyrosequencing using the forward and reverse primers in Table 1 with a KOD HotStart DNA Polymerase kit following the manufacturer’s guidelines (see Note 11). The thermal cycler parameters in this case were 40 cycles of 30 s at 95 °C, 30 s at 63 °C, and 30 s at 72 °C. 4. Create a new ‘Allele Quantification’ (AQ) assay using the Pyromark Q48 Autoprep software and type in the sequence to analyze, with the variable position designated by a slash separating the wild-type and mutated nucleotide. In our example

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Fig. 3 Heteroplasmy shift and copy number measurements in sorted cells 1 week after sorting. (a) Shift in heteroplasmy shown as difference between the condition indicated and the average value in the untreated cells. Measurements made by pyrosequencing technical PCR quadruplicates. (b) Measurement of relative mtDNA copy number by real-time PCR, normalized to the copy number of the untreated sample

case, the sequence to analyze is written as: AC/TTGTAAGACTT. Define primer addition as ‘Automatic’. 5. Set up an instrument run by loading the assay design above onto a run template. Define the number of wells to be analyzed depending on the number of PCR reactions you wish to sequence to avoid reagent waste. 6. Dilute the sequencing primer from Table 1 to 4 μM in Annealing buffer, provided with the Pyromark Advanced reagents kit. 7. Load the assay template into the Pyromark Q48 Autoprep machine and add reagents as indicated by the machine. 8. Once priming of the reagents is completed, load 3 μL of PyroMark® Q48 Magnetic beads into the number of wells required in your assay template onto a PyroMark® Q48 Disc. Add 10 μL of each PCR reaction to the corresponding well and gently pipette up and down to mix the reaction with the magnetic beads while avoiding bubbles. 9. Load disc into the machine and run pyrosequencing. The run file will give heteroplasmy values based on the relative peak intensity of the incorporated nucleotides. 3.4.2 Measuring Recovery of mtDNA Copy Number

1. For each treated sample set up 20 μL SYBR Green qPCR reactions. Use technical quadruplicates for the ND1 and HK2 genes as in Table 2. Use 10 ng of DNA per well and a primer concentration of 500 nM (see Note 12).

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2. Run the plate on a QuantStudio 3 system or equivalent system defining the run as using SYBR Green chemistry and a 20 μL reaction. 3. Export the result file and calculate the approximate copy number following the 2-ΔΔCT method [26] (see Note 13).

4

Notes 1. If selecting only the variable base does not yield high quality primer sets you may try widening the region of interest by several nucleobases on either side of the base of interest. It is crucial, however, that the variable position remains selected. 2. Unless no other assay is available, or cannot be avoided because of the nature of the mutation, it is preferable to select sequencing assays which do not dispense Adenine at the variable position. For example, if your heteroplasmic model has a variable base m.3243A > G, then it is preferable to select a primer set from the software that will measure the m.3243 T > C ratio by positioning the sequencing primer on the opposite strand. The assay design software will provide trios of primers which sequence on either the coding or complementary strand of the sequence entered, so select whichever assays are of the highest quality score and do not dispense Adenine. 3. Due to the high prevalence of NUMTs, it may be difficult to find pyrosequencing primer sets that completely avoid any amplification of nuclear DNA. Primer sets designed by the Pyromark design software with lower quality scores can be checked for nuclear off targets if the highest quality primer sets amplify several sites in nuclear DNA. 4. To ensure that the digestion of the backbones has gone to completion, also set up ligations without any of your Zinc Finger array inserts. We also recommend using a 10:1 molar ratio of zinc finger insert to backbone. 5. Considerations during clone collection: (a) If your backbone negative control culture plates contain a similar number of colonies to your zinc finger conditions, it is likely that the digestions of the backbone were inefficient and the restriction and ligation procedure may be worth repeating. (b) Eject pipette tip with picked clone directly into antibiotic containing LB broth without touching the edges of the container. (c) Give a number to each picked clone from the LB agar plate by labeling the bottom of the plate with a small circle

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and a number. This will avoid having to re-transform DH5α competent cells if the clone is successful. 6. If you are not used to gauging DNA pellet sizes, start with a smaller volume of water for resuspension, e.g., ~50 μL, as this will avoid the need for lengthy precipitation if the plasmid DNA concentration is lower than the recommended minimum of 1 μg/μL. 7. You will require 2 × 106 cells for each transformation condition, so you will need to plate an appropriate number of 175 cm2 flasks depending on the number of conditions you wish to test. 8. For two color sorting, single fluorescence controls are not required. A 5 μg mixture of both pTracer and pcmCherry is sufficient. This protocol is demonstrated with single Zinc Finger controls; however, these are not strictly required during optimization, as the FokI domains on the plasmid are obligate heterodimers. 9. The double positive cells enriched by FACS should first be tested for expression of both mtZFNs by western blotting as each construct is tagged with either the HA or FLAG epitope (Fig. 1). To make sample collection more convenient, it is recommended to split the cells into two separate wells, with one for DNA sample collection and the other one for protein. Western blotting will also conveniently allow you to verify the mitochondrial localization of each construct based on the predicted molecular weights before and after cleavage of the MTS by the mitochondrial import machinery. This procedure has been described in a previous protocol from our laboratory [17]. 10. Cells should be plated on an appropriate surface depending on the total number of double positive cells recovered from the FACS. The typical yield of double positive cells is approximately 104 cells; however, if recovered cell numbers are greater, cells can be plated on a larger surface. Useful guidelines for cell plating area are available online. To further avoid risk of contamination, the concentration of penicillin–streptomycin can be doubled to 2% of the culture medium. 11. The parameters of this PCR will require some optimization, particularly in terms of cycle number and annealing temperature, however, typically a high number of cycles (35–40) and 5 ng of DNA as the starting template are recommended. 12. Prepare Master Mixes of both Diluted DNA samples and Primers diluted in 2 × SYBR Green Master Mix: (a) For DNA: 90 ng of DNA in 81 μL total volume, which allows for 9 μL technical quadruplicates for both ND1

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and HK2 genes. Place each sample per column of a 96 well plate for convenience. Load two wells with 9 μL of the molecular grade water used to dilute your DNA samples as blanks for both ND1 and HK2 primer mixes. (b) For primers: (1.1 × total number of wells for each primer pair) μL of 10 nM F + R primer mixes in (1.1 × total number of well for each primer pair) μL of 2 × SYBR Green Master Mix, which allows for 11 μL per well of primer Master Mix. 13. Error bars can be calculated by calculating an upper and lower bound for the copy number using the following formula: Stdevtot = StdevND1 þ StdevHK2 Copy number range = 2 - ΔΔCT ± Stdevtot

Acknowledgments Medical Research Council, UK (MC_UU_00015/4) and The Lily Foundation, UK (Charity number: 1122071) are acknowledged for the support of our work. References 1. Chinnery PF, Hudson G (2013) Mitochondrial genetics. Br Med Bull 106:135–159. https://doi.org/10.1093/bmb/ldt017 2. Chacinska A, Koehler CM, Milenkovic D, Lithgow T, Pfanner N (2009) Importing mitochondrial proteins: machineries and mechanisms. Cell 138(4):628–644. https://doi.org/ 10.1016/j.cell.2009.08.005 3. Vafai SB, Mootha VK (2012) Mitochondrial disorders as windows into an ancient organelle. Nature 491(7424):7424. https://doi.org/10. 1038/nature11707 4. Wei W et al (2020) Nuclear-mitochondrial DNA segments resemble paternally inherited mitochondrial DNA in humans. Nat Commun 11(1):1740. https://doi.org/10.1038/ s41467-020-15336-3 5. Parsons TJ et al (1997) A high observed substitution rate in the human mitochondrial DNA control region. Nat Genet 15(4):4. https://doi.org/10.1038/ng0497-363 6. Lawless C, Greaves L, Reeve AK, Turnbull DM, Vincent AE (2020) The rise and rise of mitochondrial DNA mutations. Open Biol 10(5):200061. https://doi.org/10.1098/ rsob.200061

7. Elliott HR, Samuels DC, Eden JA, Relton CL, Chinnery PF (2008) Pathogenic mitochondrial DNA mutations are common in the general population. Am J Hum Genet 83(2): 254–260. https://doi.org/10.1016/j.ajhg. 2008.07.004 8. Peeva V et al (2018) Linear mitochondrial DNA is rapidly degraded by components of the replication machinery. Nat Commun 9(1): 1–11. https://doi.org/10.1038/s41467-01804131-w 9. Gammage PA, Moraes CT, Minczuk M (2018) Mitochondrial genome engineering: the revolution may not be CRISPR-Ized. Trends Genet 34(2):101–110. https://doi.org/10.1016/j. tig.2017.11.001 10. Gammage PA et al (2018) Genome editing in mitochondria corrects a pathogenic mtDNA mutation in vivo. Nat Med 24(11): 1691–1695. https://doi.org/10.1038/ s41591-018-0165-9 11. Gammage PA, Rorbach J, Vincent AI, Rebar EJ, Minczuk M (2014) Mitochondrially targeted ZFNs for selective degradation of pathogenic mitochondrial genomes bearing largescale deletions or point mutations. EMBO

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Mol Med 6(4):458–466. https://doi.org/10. 1002/emmm.201303672 12. Gammage PA et al (2016) Near-complete elimination of mutant mtDNA by iterative or dynamic dose-controlled treatment with mtZFNs. Nucl Acids Res 44(16):7804–7816. https://doi.org/10.1093/nar/gkw676 13. Minczuk M, Papworth MA, Miller JC, Murphy MP, Klug A (2008) Development of a singlechain, quasi-dimeric zinc-finger nuclease for the selective degradation of mutated human mitochondrial DNA. Nucleic Acids Res 36(12):3926–3938. https://doi.org/10. 1093/nar/gkn313 14. Doyon Y et al (2011) Enhancing zinc-fingernuclease activity with improved obligate heterodimeric architectures, Nat Meth. 8(1):1. https://doi.org/10.1038/nmeth.1539 15. Miller JC et al (2007) An improved zinc-finger nuclease architecture for highly specific genome editing. Nat Biotechnol 25(7): 778–785. https://doi.org/10.1038/nbt1319 16. Minczuk M, Papworth MA, Kolasinska P, Murphy MP, Klug A (2006) Sequence-specific modification of mitochondrial DNA using a chimeric zinc finger methylase. PNAS 103(52):19689–19694. https://doi.org/10. 1073/pnas.0609502103 17. Gammage PA, Van Haute L, Minczuk M (2016) Engineered mtZFNs for manipulation of human mitochondrial DNA heteroplasmy. In: McKenzie M (ed) Mitochondrial DNA: methods and protocols. Springer, New York, pp 145–162. https://doi.org/10.1007/978-1-4939-30401_11 18. Bhakta MS, Segal DJ (2010) The generation of zinc finger proteins by modular assembly. In: Mackay JP, Segal DJ (eds) Engineered zinc finger proteins: methods and protocols. Humana Press, Totowa, pp 3–30. https://doi. org/10.1007/978-1-60761-753-2_1

19. Wright DA et al (2006) Standardized reagents and protocols for engineering zinc finger nucleases by modular assembly. Nat Protoc 1(3):1637–1652. https://doi.org/10.1038/ nprot.2006.259 20. Sander JD et al (2011) Selection-free zincfinger-nuclease engineering by contextdependent assembly (CoDA). Nat Meth 8(1): 1. https://doi.org/10.1038/nmeth.1542 21. Maeder ML, Thibodeau-Beganny S, Sander JD, Voytas DF, Joung JK (2009) Oligomerized pool engineering (OPEN): an ‘open-source’ protocol for making customized zinc-finger arrays. Nat Protoc 4(10):1471–1501. https:// doi.org/10.1038/nprot.2009.98 22. Isalan M (2021) DNA recognition/processing|zinc fingers: structure and design☆. In: Jez J (ed) Encyclopedia of biological chemistry III, 3rd edn. Elsevier, Oxford, pp 506–516. https://d oi.org/10.1016 /B97 8-0-12809633-8.21266-1 23. Klug A (2010) The discovery of zinc fingers and their applications in gene regulation and genome manipulation. Annu Rev Biochem 79(1):213–231. https://doi.org/10.1146/ annurev-biochem-010909-095056 24. Kauppila JHK et al (2016) A phenotype-driven approach to generate mouse models with pathogenic mtDNA mutations causing mitochondrial disease. Cell Rep 16(11):2980–2990. https://doi.org/10.1016/j.celrep.2016. 08.037 25. “Nucleotide BLAST: Search nucleotide databases using a nucleotide query.” https://blast. ncbi.nlm.nih.gov/Blast.cgi?PAGE_TYPE= BlastSearch. Accessed 19 May 2021 26. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C (T)) method. Methods 25(4):402–408. https://doi.org/10.1006/meth.2001.1262

Chapter 24 Biolistic Transformation of Chlamydomonas reinhardtii and Saccharomyces cerevisiae Mitochondria Nathalie Bonnefoy and Claire Remacle Abstract Chlamydomonas reinhardtii and Saccharomyces cerevisiae are currently the two micro-organisms in which genetic transformation of mitochondria is routinely performed. The generation of a large variety of defined alterations as well as the insertion of ectopic genes in the mitochondrial genome (mtDNA) are possible, especially in yeast. Biolistic transformation of mitochondria is achieved through the bombardment of microprojectiles coated with DNA, which can be incorporated into mtDNA thanks to the highly efficient homologous recombination machinery present in S. cerevisiae and C. reinhardtii organelles. Despite a low frequency of transformation, the isolation of transformants in yeast is relatively quick and easy, since several natural or artificial selectable markers are available, while the selection in C. reinhardtii remains long and awaits new markers. Here, we describe the materials and techniques used to perform biolistic transformation, in order to mutagenize endogenous mitochondrial genes or insert novel markers into mtDNA. Although alternative strategies to edit mtDNA are being set up, so far, insertion of ectopic genes relies on the biolistic transformation techniques. Key words Mitochondrial DNA, Genetic transformation, Biolistic techniques, Homologous recombination, Mutagenesis and ectopic gene insertion, Chlamydomonas reinhardtii, Saccharomyces cerevisiae

1

Introduction Transformation of the mitochondrial genome can be achieved in two unicellular micro-organisms: originally the yeast Saccharomyces cerevisiae and later on the green microalga Chlamydomonas reinhardtii [1]. In both systems, donor DNA is delivered into mitochondria by microprojectile bombardment (biolistic transformation) and the DNA is inserted into the recipient mitochondrial genome by homologous recombination. The use of a biolistic particle delivery

Supplementary Information The online version contains supplementary material available at https://doi.org/ 10.1007/978-1-0716-2922-2_24. Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_24, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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system where thousands of microprojectiles are coated with multiples copies of the donor DNA is an efficient way to target the multicopy mitochondrial genome, since it increases the chances to deliver enough copies of the donor DNA to invade the endogenous polyploid genome. In addition, the use of adequately sized microparticles and a rupture disk determining the power of the helium shockwave allows the particles to cross both the cell and mitochondrial membranes [2]. In other organisms, mitochondrial transformation has been unsuccessful, but DNA editing of the mitochondrial genome is becoming feasible in both animals [3] and plants [4], using sequence-specific nucleases targeted to mitochondria. However, so far, insertion of novel genes into the mitochondrial genome is still only possible through biolistic transformation, i.e., in S. cerevisiae and C. reinhardtii, and should potentially be achievable in Candida glabrata, where biolistic transformation has also been reported [5]. The first demonstration of mitochondrial transformation in Chlamydomonas was published in 1993 [6]. The authors used a biolistic delivery system to transform a mutant called dum1 that harbors a 1.5 kb terminal deletion of the linear 15.8 kb mitochondrial genome comprising the telomere (500 bp) and most of the cob gene (1 kb) encoding apocytochrome b (Fig. 1). In addition to this deleted genome, the mutant also contains dimers which result from head-to-head fusions between deleted monomers [6, 7]. The dum1 mutant is unable to grow in the dark in the presence of acetate because of the loss of the activity of complex III and is a non-reverting mutant, which after selection in the dark allows the recovery of rare respiratory competent cells with intact mtDNA. The next step in the setup of mitochondrial transformation was reached in 2005 when Yamasaki and coauthors [8] reported that

1.1 Chlamydomonas Mitochondrial Transformation

cob

nd4

dum22 dum1 dum11

nd5

cox1

nd2

W nd6

Q

nd1

L6 L5 L7 S1 S2

M

rtl

L3

L8

L4 S3

L1 L2 S4

Fig. 1 Physical map of the C. reinhardtii 15.8 kb linear mitochondrial genome (GenBank accession EU306622). Horizontal blue arrows indicate the bidirectional transcription origin. The blue boxes represent the eight protein-coding genes: cob (apocytochrome b of complex III), cox1 (subunit 1 of cytochrome c oxidase or complex IV), nd1, 2, 4, 5, and 6 (subunits of NADH:ubiquinone oxidoreductase or complex I) and rtl (reverse transcriptase-like protein). The boxes in two shades of green represent LSU (L1–L8) and SSU (S1–S4) genes encoding the structural ribosomal RNA fragments. The vertical bars correspond to the 3 tRNA genes with the one-letter code (W, Q, and M). The light blue arrows at each end of the genome represent the inverted terminal repeats

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Table 1 Experimental conditions for mitochondrial transformation in C. reinhardtii Transformation ratea References

Strain

Beads

Exogenous DNA

dum1

Tungsten beads (~1 μM)

Partially purified C. reinhardtii mitochondrial genome (0.8 μg per bombardment)

3.5–12.5b

[6]

dum1, dum14, dum16

Gold beads (600 + 100 nm)c

Plasmids containing 1.8–5.0 kb mtDNA fragments (5 μg per bombardment) 3.8 kb PCR producte (5 μg per bombardment)

5.5b,d

[8]

Plasmids containing 1.8–6.6 kb mtDNA fragments (3 μg per bombardment) PCR product (1.5 μg per bombardment)

~100–220f (10–22b)

dum11

Tungsten beads ( /output/Sample.sam

4. Sort and index the Sam file using Picard 2.4.1 to generate a . bam file with the following command lines: java -Xmx4G -jar picard.jar SortSam INPUT = Sample.sam O=Sample_sorted.bam SORT_ORDER = coordinate; java -Xmx4G -jar picard.jar BuildBamIndex INPUT = Sample_sorted.bam

5. Use the sorted bam file as the input for Mutserve software. There are two option for its usage: either upload the sorted bam file online at the Mitoverse website using the mtDNAServer v2 tool; or, download the Haplocheck suite and launch the following command (see Note 12): java -Xmx4G -jar mutserve.jar call --level 0.01 --reference / reference_mtDNA_path/chrM.fasta --mapQ 20 --baseQ 20 --deletions --output /output/variants.vcf.gz --no-ansi Sample_sorted.bam --threads 4 --write-raw

3.5 Output Visualization and Interpretation

1. Open the generated sorted bam and vcf.gz files using Integrative Genomics Viewer (IGV) tool. This tool allows visualization of aligned reads, depth of coverage profile along the mitochondrial genome, and homoplasmic/heteroplasmic variants (Fig. 2). To correctly open the bam and vcf files, load the reference fasta file used for aligning the reads as a reference

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A

Depth of Depth of coverage coverage

Sample1 Sample2 Sample3

B

m.13506T>C Cov = 1669x T=0%, C=100%

m.13513G>A Cov = 1686x G=34%, A=66%

Fig. 2 NGS data visualization with the integrative genomics viewer (IGV) tool. (a) Coverage profiles of the sorted bam files from 3 mtDNA sequencing, visualized using IGV tool. The height (y-axis value) of the signal corresponds to the depth of coverage. Gray bars correspond to reference nucleotides, color bars correspond to alternative nucleotides (C: blue, A: green, G: yellow; T: red). (b) Zoomed-in visualization of aligned reads, showing a homoplasmic nucleotide change (m.13506T>C, belonging to haplogroup L) and a heteroplasmic nucleotide change (m.13513G>A, pathogenic variant associated with Leigh syndrome). For each mtDNA position, it is possible to visualize the depth of coverage, and percentages (i.e., heteroplasmy level) of each nucleotide

genome file in IGV tool. Variants will be color coded, and to visualize their statistics (i.e., heteroplasmic percentage) click on the variant position. 2. The output generated by Mutserve/Mitoserve will consist of a variants.txt or variants.vcf.gz file with a list of homoplasmic and/or heteroplasmic variants (Table 1), and an annotated variants_ann.txt file (Table 2) with information associated with the sequenced variants, such as their position, the reference and the alternative base, coverage statistics and heteroplasmic percentages, gene names and amino acid changes.

G

C

A

G

PASS 11098 A

PASS 12933 A

Sample.bam PASS 13513 G

PASS 13759 G

PASS 14587 A

PASS 15326 A

PASS 16092 T

PASS 16256 C

PASS 16293 A

PASS 16301 C

PASS 16311 T

Sample.bam

Sample.bam PASS 13506 T

PASS 13676 A

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

Sample.bam

100%

100%

100%

1%

100%

100%

99%

99%

1%

66%

100%

1%

1%

1%

C

T

G

C

C

G

G

A

A

A

C

A

A

T

0%

0%

0%

99%

0%

0%

0%

0%

99%

66%

0%

99%

99%

99%

0% 0%

– –

0%

–

0%

0%

–

–

0%

–

1%

0%

–

A

1%

G

34%

0%

– G

1%

1%

1%

G

G

C

6038

6982

6705

7911

5394

2243

1865

915

637

1641

1640

982

1937

1120

2340

2525

2287

2984

2459

1045

1053

547

430

560

560

280

1254

499

3698

4457

4418

4927

2935

1198

812

368

207

1081

1080

702

683

621

Pos position in the mtDNA, Ref reference nucleotide according to NC_012920, FWD forward reads, REV reverse reads. D: single-nucleotide deletion. The variants depicted in Fig. 2b are reported in bold

C

T

G

A

C

G

G

A

G

C

PASS 10477 T

Sample.bam

Ref Variant VariantLevel MajorBase MajorLevel MinorBase MinorLevel Coverage CoverageFWD CoverageREV

Filter Pos

ID

Table 1 Excerpt from the Variant Calling file by Mitoverse

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Table 2 Excerpt from the Variant Annote file by Mitoverse, reporting annotations for the variant m.13513G>A ID

Mutation Substitution Maplocus Category AAC

Sample 13513A Transition Name

MTND5

MutPred_ OXPHOS rCRS_ CI_MitoTool complex NuMTs_dayama Surr_seq Score

Coding D393N 0.834

1

I

0

CCAAA [G/A] ACCAC

AAC amino acid change, CI conservation index by Mitotool, OXPHOS oxidative phosphorylation, NuMTs nuclear mitochondrial insertion sequence by Dayama et al. 2014 [27], rCRS revised Cambridge Reference Sequence, Surr_seq surronding sequence

4

Notes 1. DNA can be extracted from different biological specimens. We routinely sequence mtDNA starting from total DNA extracted from: blood, fibroblast cell culture, muscle tissue, saliva or buccal swab, and epithelial urinary cells (UEC). Heteroplasmy levels of the same mtDNA variant can vary across tissues from the same individual, with blood usually carrying the lowest levels. For this reason, selection of the most appropriate tissue for mtDNA extraction has to be taken into serious account before proceeding with NGS analysis. 2. To check the heteroplasmy levels of an already known mutation (e.g., to study different tissues or for segregation analysis in family members), the same protocol described above for one amplicon long range PCR can be applied to short targeted PCR-amplicons spanning the region containing the mtDNA variant [20]. 3. With the proposed primers (Subheading 2.1, items 9–10), ~50 nucleotides corresponding to the primer annealing region (m.16401–16449) cannot be evaluated for mutations. Notably, this region is located in the non-coding D-loop region and no pathogenic variants are known to be located here. 4. Pre-heating of the thermal cycler to 93 °C is essential for increasing the yield of the desired amplified product (Subheading 3.1, step 9). 5. Gel electrophoresis analysis of the PCR product (Subheading 3.1, step 8) should show a unique band at a molecular weight of 16.5 kb; however, in the case of large deletions affecting mtDNA, it may be possible to notice one or more additional bands at lower molecular weight (Fig. 1). 6. If no bands are present in the gel (Subheading 3.1, step 8) (Fig. 1), possible explanations include: (i) DNA degradation that may impede the correct synthesis of the large amplicon, or

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(ii) a mtDNA variant localized within the primer annealing region (m.16401–16449). In the first case, amplification of shorter overlapping amplicons is suggested [20]. In the second scenario, the use of an alternative couple of primers could solve the issue (for instance, we used 5681-Fw 5′-caaacacttagttaacagctaagca-3′ and 5680-Rw 5′-tgggtttaagtcccattggt-3′). 7. For the Nextera XT DNA Library Preparation (Subheading 3.3, step 3), multiplexed sample processing is possible thanks to the use of up to 96 different barcoding indexes. This step, which allows large numbers of samples to be pooled and handled/processed simultaneously, is recommended in order to increase throughput and save time (a detailed guide for sample multiplexing and indexes selection can be found in Illumina Document #1000000041074). 8. Since the mitochondrial genome length is just 16.5 kb, the sequencing of one or a few mtDNA samples (Subheading 3.3, step 3) on a MiSeq run will generate an extraordinary number of reads per sample, exceeding the need for a proper depth of coverage by far. For this reason, we suggest either large numbers of samples (up to 96) or combining mtDNA samples prepared with the Nextera XT DNA Library Prep Kit with samples processed with other Nextera applications, such as targeted gene panels. In the latter case, it is important to correctly balance the two types of libraries depending on the total length of sequenced regions and on samples number. 9. The quality control check (Subheading 3.4, step 1) allows determination of the percentage of reads suitable for all the following bioinformatics steps (alignment, variant calling etc.): a good run leads to 85–90% reads passing this control. 10. In the trimming step (Subheading 3.4, step 2), the parameter MINLEN:100 means that reads shorter than 100 bp are excluded from the alignment. 100 is the value suggested when using a read length of 201 bp (Subheading 3.3, step 4); in case of shorter reads set up in the sample sheet, the MINLEN parameter can be decreased. 11. Alignment of the reads produced by NGS is performed by BWA (Subheading 3.4, step 3), a widely diffused tool for the alignment of short reads. This tool is not developed for the identification of large deletions; other aligners have been specifically developed for this kind of purpose (e.g., BBMap). 12. The proposed bioinformatics pipeline (Subheading 3.4) allows processing of raw fastq files (fastq is the standard output file format generated by NGS sequencing platforms). However, the MiSeq Sequencing Systems is able to generate fastq files already trimmed and ready to be aligned to the mtDNA

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reference sequence (Subheading 3.4, step 3). MiSeq can also generate bam files. These files can be visualized by IGV, but, since the MiSeq system uses the whole human genome as a reference sequence (hg19), the generated aligned files cannot be used as input for Mutserve (Subheading 3.5, step 1). This tool accepts only bam files aligned to the reference rCRS mtDNA sequence.

Acknowledgments Our work was supported by Associazione Luigi Comini Onlus (CV), Telethon (GGP20013 to CV), Starting grant from Department of Biomedical Sciences of University of Padova (CV), the Italian Ministry of Health (RCC funding to AL and DG), and the Pierfranco e Luisa Mariani Foundation (AL, DG). References 1. Gorman GS, Chinnery PF, DiMauro S et al (2016) Mitochondrial diseases. Nat Rev Dis Primers 2:16080. https://doi.org/10.1038/ nrdp.2016.80 2. Nunnari J, Suomalainen A (2012) Mitochondria: in sickness and in health. Cell 148:1145– 1159. https://doi.org/10.1016/j.cell.2012. 02.035 3. Wallace DC, Chalkia D (2013) Mitochondrial DNA genetics and the heteroplasmy conundrum in evolution and disease. Cold Spring Harb Perspect Biol 5:a021220. https://doi. org/10.1101/cshperspect.a021220 4. Gasparre G (2020) The human mitochondrial genome. Elsevier, MA 5. Chinnery PF, Gomez-Duran A (2018) Oldies but goldies mtDNA population variants and neurodegenerative diseases. Front Neurosci 12:682. https://doi.org/10.3389/fnins. 2018.00682 6. Amorim A, Fernandes T, Taveira N (2019) Mitochondrial DNA in human identification: a review. PeerJ 7:e7314. https://doi.org/10. 7717/peerj.7314 7. Schlieben LD, Prokisch H (2020) The dimensions of primary mitochondrial disorders. Front Cell Dev Biol 8:600079. https://doi. org/10.3389/fcell.2020.600079 8. Holt IJ, Harding AE, Morgan-Hughes JA (1988) Deletions of muscle mitochondrial DNA in patients with mitochondrial myopathies. Nature 331:717–719. https://doi.org/ 10.1038/331717a0

9. Wallace DC, Singh G, Lott MT et al (1988) Mitochondrial DNA mutation associated with Leber’s hereditary optic neuropathy. Science 242:1427–1430. https://doi.org/10.1126/ science.3201231 10. Zeviani M, Moraes CT, DiMauro S et al (1988) Deletions of mitochondrial DNA in KearnsSayre syndrome. Neurology 38:1339–1346. https://doi.org/10.1212/wnl.38.9.1339 11. Luft R, Ikkos D, Palmieri G et al (1962) A case of severe hypermetabolism of nonthyroid origin with a defect in the maintenance of mitochondrial respiratory control: a correlated clinical, biochemical, and morphological study. J Clin Invest 41:1776–1804. https:// doi.org/10.1172/JCI104637 12. Brown MD, Torroni A, Reckord CL, Wallace DC (1995) Phylogenetic analysis of Leber’s hereditary optic neuropathy mitochondrial DNA’s indicates multiple independent occurrences of the common mutations. Hum Mutat 6:311–325. https://doi.org/10.1002/humu. 1380060405 13. Goto Y, Nonaka I, Horai S (1990) A mutation in the tRNA(Leu)(UUR) gene associated with the MELAS subgroup of mitochondrial encephalomyopathies. Nature 348:651–653. https://doi.org/10.1038/348651a0 14. Wallace DC, Zheng XX, Lott MT et al (1988) Familial mitochondrial encephalomyopathy (MERRF): genetic, pathophysiological, and biochemical characterization of a mitochondrial DNA disease. Cell 55:601–610. https:// doi.org/10.1016/0092-8674(88)90218-8

mtDNA Sequencing and Heteroplasmy Quantification by NGS 15. Holt IJ, Harding AE, Petty RK, MorganHughes JA (1990) A new mitochondrial disease associated with mitochondrial DNA heteroplasmy. Am J Hum Genet 46:428–433 16. Zeviani M (2004) Mitochondrial disorders. Brain 127:2153–2172. https://doi.org/10. 1093/brain/awh259 17. Payne BAI, Wilson IJ, Yu-Wai-Man P et al (2013) Universal heteroplasmy of human mitochondrial DNA. Hum Mol Genet 22: 384–390. https://doi.org/10.1093/hmg/ dds435 18. Lyons EA, Scheible MK, Sturk-Andreaggi K et al (2013) A high-throughput Sanger strategy for human mitochondrial genome sequencing. BMC Genomics 14:881. https://doi.org/10. 1186/1471-2164-14-881 19. Yan J, Zhang R, Xiong C et al (2014) Pyrosequencing is an accurate and reliable method for the analysis of heteroplasmy of the A3243G mutation in patients with mitochondrial diabetes. J Mol Diagn 16:431–439. https://doi. org/10.1016/j.jmoldx.2014.03.005 20. Legati A, Zanetti N, Nasca A et al (2021) Current and new next-generation sequencing approaches to study mitochondrial DNA. J Mol Diagn 23:732. https://doi.org/10. 1016/j.jmoldx.2021.03.002 21. Zhang W, Cui H, Wong L-JC (2012) Comprehensive one-step molecular analyses of mitochondrial genome by massively parallel sequencing. Clin Chem 58:1322–1331. https://doi.org/10.1373/clinchem.2011. 181438

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22. Levy MA, Kerkhof J, Belmonte FR et al (2021) Validation and clinical performance of a combined nuclear-mitochondrial next-generation sequencing and copy number variant analysis panel in a Canadian population. Am J Med Genet 185:486–499. https://doi.org/10. 1002/ajmg.a.61998 23. Falk MJ, Pierce EA, Consugar M et al (2012) Mitochondrial disease genetic diagnostics: optimized whole-exome analysis for all MitoCarta nuclear genes and the mitochondrial genome. Discov Med 14:389–399 24. Wagner M, Berutti R, Lorenz-Depiereux B et al (2019) Mitochondrial DNA mutation analysis from exome sequencing-A more holistic approach in diagnostics of suspected mitochondrial disease. J Inherit Metab Dis 42:909– 917. https://doi.org/10.1002/jimd.12109 25. Hazkani-Covo E, Zeller RM, Martin W (2010) Molecular poltergeists: mitochondrial DNA copies (numts) in sequenced nuclear genomes. PLoS Genet 6:e1000834. https://doi.org/10. 1371/journal.pgen.1000834 26. Li M, Schroeder R, Ko A, Stoneking M (2012) Fidelity of capture-enrichment for mtDNA genome sequencing: influence of NUMTs. Nucleic Acids Res 40:e137. https://doi.org/ 10.1093/nar/gks499 27. Dayama G, Emery SB, Kidd JM, Mills RE (2014) The genomic landscape of polymorphic human nuclear mitochondrial insertions. Nucleic Acids Res 42(20):12640–12649. https://doi.org/10.1093/nar/gku1038

Chapter 27 Genomic Strategies in Mitochondrial Diagnostics Dasha Deen, Charlotte L. Alston, Gavin Hudson, Robert W. Taylor, and Angela Pyle Abstract Pathogenic variants in both mitochondrial and nuclear genes contribute to the clinical and genetic heterogeneity of mitochondrial diseases. There are now pathogenic variants in over 300 nuclear genes linked to human mitochondrial diseases. Nonetheless, diagnosing mitochondrial disease with a genetic outcome remains challenging. However, there are now many strategies that help us to pinpoint causative variants in patients with mitochondrial disease. This chapter describes some of the approaches and recent advancements in gene/variant prioritization using whole-exome sequencing (WES). Key words Genetic diagnosis, Genomics, Mitochondrial disease, Whole-exome sequencing, Variant detection, Variant annotation, Clinical reporting

1

Introduction

1.1 Genomic Strategies for Mitochondrial Disease Diagnostics

Genomic strategies that are well established for the detection of causal variants of mitochondrial diseases include candidate gene sequencing, sequencing of specific gene panels, whole-exome sequencing (WES) and whole genome sequencing (WGS) [1, 2] (Table 1). Alternative approaches such as transcriptome sequencing and metabolome and proteome analysis are increasingly being used to supplement next-generation sequencing (NGS) and improve diagnosis rates [3–5]. When a patient presents with a mitochondrial phenotype, mutations in mitochondrial DNA (mtDNA) are first excluded, either via Sanger sequencing or using NGS technologies [6]. If mtDNA sequencing fails to identify a causative variant, then the patients’ nuclear DNA (nDNA) is investigated. If there is an appropriate gene panel available, typically dictated by phenotype, for

Supplementary Information The online version contains supplementary material available at https://doi.org/ 10.1007/978-1-0716-2922-2_27. Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_27, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Table 1 Pros and Cons of different genomic strategies for mitochondrial disease diagnostics Candidate gene Sanger Genomic strategy sequencing

Gene panel sequencing

Whole-exome sequencing

Target

1–5 genes (typically only coding exons)

40–400 genes

22,000 genes

All genes, non-coding DNA and translocations

Types of variants detected

SNVs, small indels

SNVs, indels, CNVs

SNVs, indels, CNVs

SNV, indels, CNVs, structural variants

Ease of computational analysis

Easy

Easy

Easy

Requires significant computational power; reliably detecting structural variants might be a challenge

Ease of interpretation

Straightforward Straightforward Straightforward Few in silico resources are available for structural variants and variants in noncoding regions pathogenicity prediction

Examples of publications

[44]

[45]

[46]

Whole genome sequencing

[47]

example, when isolated mitochondrial complex I deficiency is observed, then the patient nDNA will be sequenced for targeted genes. Examples where such approach is appropriate are further discussed in [3]. However, due to the heterogeneity of mitochondrial disease, the use of gene panels might not provide a high diagnostic yield [2]. WES, the focus of this chapter, is currently the most widely used approach for mitochondrial diagnostics due to the balance between the cost, amount of information obtained, and ease of computational analysis [3]. However, this is likely to change given the year-on-year reduction in the cost of whole genome sequencing (WGS) [7]. Furthermore, in addition to detecting single nucleotide variants (SNVs) and small indels (2–50 bp), WGS also allows more robust detection of structural and copy number variants, repeat variants, noncoding SNVs and improved detection of larger indels (>50 bp) [8], although diagnostic strategies for prioritization of these types of variants remain insufficiently developed. Overall, to be able to detect the causative variant of a mitochondrial disease using any genomic strategy mentioned above, three conditions should be met:

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• The appropriate gene set must be selected for the analysis (i.e., the variant is in a gene that was included in the gene panel). • The platform and software tools used are adequate for detecting the type of variants. • Pathogenic variants in the gene/noncoding regions are known to be associated with mitochondrial disease. It is worth reanalysing unsolved diagnostics cases as the knowledge about the genes and variants associated with mitochondrial diseases keeps expanding, and software tools become more refined. 1.2 Sample Selection Strategies

The most successful approach for mitochondrial diagnostics is sequencing multiple individuals, particularly affected individuals and their parents (case-parent trio studies). This strategy allows investigators to incorporate information regarding the mode of inheritance and detect causative rare variants, including de novo variants. However, obtaining multiple samples is not always possible due to the cost of analysis and ethical constraints. One replicate is enough for each patient. Blood, muscle or fibroblasts can be used as a source of genomic DNA [9]. Some pathogenic mtDNA variants are restricted to skeletal muscle due to mosaicism, meaning that analysis of blood or another non-invasive tissue would be uninformative for that individual. Moreover, while perhaps present at detectable levels in early life, some mtDNA variants undergo negative selection, and, therefore, may evade detection in blood if sampling was performed in later life [10]. For Mendelian, nuclear genetic variants, blood would be an entirely appropriate tissue. Often an approach is used where a single individual is analyzed, and then, the prioritized variants are checked individually for the mode of inheritance. When running a variant detection workflow on a single patient sample, where allelic segregation within a family cannot be checked, the investigator may inevitably identify a large list of potentially causative variants. This list can include common artefacts that arise during sample preparation, sequencing and variant calling, as well as variants that are rare overall but are common in a population, where data on allele population frequency are unavailable. The impact of sequencing artefacts can be mitigated, somewhat, by sequencing and analysing several (unrelated) samples together to identify run or batch-specific anomalous variant calls.

1.3 Variant Detection and Characterization Workflow

Clinical variant detection workflows begin with the patient’s DNA/tissue sample and DNA extraction, and then follow a typical pathway of sequencing, quality control, read alignment, variant calling, variant annotation, variant prioritization, and clinical report generation (Fig. 1). In many cases, these workflows are designed, maintained and operated by dedicated bioinformatics personnel. As bioinformatics software is modular in nature, each computational step can be achieved using different tools (Table 2). Tools

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Fig. 1 Workflow showing steps and tools required for the diagnostic genomic testing of patients with suspected mitochondrial disease Table 2 Selected list of whole exome analysis tools for pre-processing, read alignment, variant calling and variant annotation Workflow

Category

Software tools

Preprocessing

Raw data QC

FastQC

Read trimming

Trimmomatic

Read alignment

BWA

Read alignment and processing

Bowtie Read duplication removal

Picard Samtools Sambamba

Variant Detection (SNVs and indels) and annotation

Variant calling

GATK Platypus VarScan2 DeepVariant

Variant annotation

VEP AnnoVar Jannovar

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for the initial data processing are relatively mature and generally produce similar results, while many options exist for variant detection algorithms [see [11] for a recent review]. No single variant caller for any variant type performs optimally under all conditions (see Table 2 for the commonly used variant callers). GATK HaplotypeCaller is being actively developed by the Broad Institute [12] and appears to be the most widely adopted tool in clinical variant detection pipelines for detecting SNVs and small indels ( fastq/subject_R1.fastq.gz cat subject1_L001_R2_001.fastq.gz \ subject1_L002_R2_001.fastq.gz > fastq/subject_R2.fastq.gz

5. Check the quality of the sequenced data files using FastQC software. fastqc fastq/subject_R1.fastq.gz --outdir=qc fastqc fastq/subject_R1.fastq.gz --outdir=qc

FastQC will produce an html format report about the sequencing data quality in the ‘qc’ folder. The main characteristics to look at are the sequence quality scores, which are measured in Phred quality scores: Q = - log 10 E where E is the probability of the error. A higher Phred score indicates a higher probability that a sequenced base is called correctly.

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Characteristics to look at are (see Note 6): – The per base sequence quality scores along the whole length of the read should not generally drop below Phred score 30, which gives 99.9% accuracy in the sequenced base. However, a deterioration of the quality of sequencing sometimes happens towards the end of the read. – The average quality score per sequence should also be higher than Phred Score 30. – The plot of per sequence GC for a whole-exome sequencing run should be unimodal; the location of the peak relative to % GC content depends on the kit used. – There should be no overrepresented sequences detected for WES data sets. Contact the sequencing facility if there are overrepresented sequences present as this might indicate the problems with the sequencing library preparation and might affect overall coverage. 6. Optional: check for sample contamination with DNA from other organisms using FastQ screen [24]: fastq_screen fastq/subject1_R1.fastq.gz --aligner bwa --force --outdir qc fastq_screen fastq/subject1_R2.fastq.gz --aligner bwa –force --outdir qc

FastQ screen takes a subset of reads from sample *.fastq files and tries to align it to the genomes of model organisms as well as common contamination sequences. The FastQ screen will produce a *.txt file describing how many reads were aligned to the respective genomes. The majority of the reads (at least 80%) should be aligned to the human genome, although 10–15% of reads are expected to align to the mouse and rat genomes because of regions homologous to human genomes present there. This proportion will be higher for shorter reads and single-end reads. A cause for concern should be a significant proportion of reads (more than 5%) aligned to the genomes of other model organisms other than human, mouse or rat or of unknown origin. Unknown origin usually indicates that the genome of the species with which the sample is contaminated is not in the list of the genomes used by FastQ screen. 7. Make an index for the reference genome: bwa index ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ ref/GCA_000001405.15_GRCh38_no_alt

This step will need to be done only once to generate the index.

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8. Align the pair-end DNA sequencing reads to the reference human genome using bwa and convert SAM files to BAM files using samtools [25]: bwa mem ref/GCA_000001405.15_GRCh38_no_alt fastq/subject1_R1.fastq.gz \ fastq/subject1_R2.fastq.gz -M \ -R "@RG\tID:FlowCell.subject1\tSM:subject1\tPL:illumina\tLB:mito.subject1" | \ samtools sort - -O bam | tee bam/subject1_bwa_output.bam | \ samtools index - bam/subject1_bwa_output.bam.bai

This is the most computationally demanding step, and it can take a couple of hours to align the sequenced reads to the genome. The alignment is recommended to run with the default parameters as increasing the sensitivity of the alignment might lead to the higher level of false positive variants in downstream steps. Alternative step 8: If the computer is at the lower end of the recommended specification, separate the steps of aligning and converting the files from sam to bam. However, the sam files generated by bwa aligner will be large (tens of GBs), so enough free space should be available on the hard drive. Sam files can be deleted after they are converted into bam files. bwa mem ref/GCA_000001405.15_GRCh38_no_alt fastq/subject1_R1.fastq.gz fastq/ \ subject1_R2.fastq.gz -M \ -R "@RG\tID:FlowCell. subject1\tSM: subject1\tPL:illumina\tLB:mito.subject1" > \ bam/subject1.sam samtools sort -O bam bam/subject1.sam > bam/subject1.bam samtools index bam/subject1.bam

9. Remove duplicated reads that originate from a single fragment of DNA using Picard Tools and index the resulting deduplicated bam file: java -jar picard.jar MarkDuplicates \ I=bam/subject1.bam \ O=bam/subject1_dedup.bam \ REMOVE_DUPLICATES=true \ M=qc/subject1_dup_metrics.txt samtools index bam/subject1_dedup.bam

10. Check the quality of the alignments: (a) calculate read alignment statistics: samtools flagstat bam/subject1_dedup.bam > qc/subject1_stat.txt

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The majority of the reads (at least 95%) should be aligned to the human genome. Lower alignment rates indicate either low sequence quality of the reads, contamination of DNA samples with technical sequences/DNA from other organisms or errors in the alignment process itself. (b) (optional) calculate the coverage: First, convert the bed file with the exons expected to be covered by the exome capture kit being used into IntervalList: java -jar picard.jar BedToIntervalList \ I=ref/exons_covered.bed \ O=ref/exons_covered.interval_list \ SD=ref/GCA_000001405.15_GRCh38_no_alt

This command needs to be run only once to generate an IntervalList format. Then, calculate the coverage: java -jar picard.jar CollectWgsMetrics \ I=bam/subject1_dedup.bam \ O=qc/subject1_coverage.txt \ R= ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ COUNT_UNPAIRED=true \ INTERVALS=ref/exons_covered.interval_list

Coverage depicts what proportion of the exome was sequenced and at what depth. The regions of the genome with missing coverage will be unavailable for variant detection. The regions of the genome with low coverage will exhibit lower sensitivity and specificity in variant detection. It is important to make sure that the regions of interest (e.g., exons of Mitocarta genes, exons of likely causative genes) have sufficient and even coverage (see Note 3). (c) (optional) check whether the sex of the sample matches the sex in the records of the patients samtools idxstats bam/subject1_dedup.bam > qc/subject1_idxstat.txt

This command will display how many reads map to different chromosomes. For a sample from a male around 75–80% of total reads mapping to sex chromosomes will be mapped to the X chromosome, while only 20–25% will be mapped to the Y chromosome due to differences in X and Y chromosome sizes. For a sample from a female up to

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5% of total reads mapping to sex chromosomes will be mapped to the Y chromosome, due to the Y chromosome containing regions of homology with autosomes. (d) (optional) check whether the sample has been contaminated by other human samples: VerifyBamID \ --BamFile bam/subject1_dedup.bam \ --SVDPrefix hgdp.100k.b38.vcf.gz.dat \ --Reference ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ --Output qc/subject1

verifyBamID software [26] checks whether the reads are contaminated as a mixture of two samples by calculating expected allele frequencies. If the percentage of sample contamination detected is above 2%, the sample should be investigated further to identify the possible contamination, and the results from this sample might be unreliable. 11. (optional) Combine quality control metrics in a single html report: multiqc qc -o final_output -n QC_report

12. Call variants using GATK HaplotypeCaller, separate for each sample: gatk --java-options "-Xmx40g" HaplotypeCaller \ -R ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ -I bam/subject1_dedup.bam \ -O vcf/subject1.g.vcf.gz \ -ERC GVCF \ --do-not-run-physical-phasing true

This step will generate gVCF files (g.vcf.gz extension), which store the sequencing information for every position in the genome, both variant and non-variant. This step is computationally demanding. 13. If several samples are run at the same time, combine the gVCF files with the variants from different samples in a single file: gatk CombineGVCFs \ -R ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ --variant vcf/subject1.g.vcf.gz \ --variant vcf/subject2.g.vcf.gz \ --variant vcf/subject3.g.vcf.gz \ -O vcf/final.g.vcf.gz

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In this example, gVCF files from three samples are combined. Combining the samples from the same run and/or pedigree in a single gVCF file will increase accuracy of variant calling. Alternatively, if only one sample is run through the workflow, index the gVCF file for the sample; here, the first step is renaming the file to keep it consistent with the rest of the chapter: mv vcf/subject1.g.vcf.gz vcf/final.g.vcf.gz gatk IndexFeatureFile -I vcf/final.g.vcf.gz

14. Call the genotypes of the samples: gatk --java-options "-Xmx40g" GenotypeGVCFs \ -R ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ -V vcf/final.g.vcf.gz \ -O vcf/final.vcf.gz

This step will output a vcf file that stores the sequencing information only for the positions in the genome that are different from the reference and will try to assign HET or HOM genotypes to it. The step is computationally demanding. 15. Filter variants based on sequence quality. This step removes the variants that do not pass the quality thresholds and are likely to be artefacts. (a) First separate the called variants into SNPs and indels, since the quality parameters are different for each: gatk SelectVariants \ -V vcf/final.vcf.gz \ -select-type SNP \ -O vcf/final_snps.vcf.gz gatk SelectVariants \ -V vcf/final.vcf.gz \ -select-type INDEL \ -O vcf/final_indels.vcf.gz

(b) Filter SNP and indel variants based on quality: gatk VariantFiltration \ -V vcf/final_snps.vcf.gz \ -filter "QD < 2.0" --filter-name "QD2" \ -filter "QUAL < 30.0" --filter-name "QUAL30" \

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GATK offers two ways of filtering the quality of the variants: using predefined parameters (hard filtering) and using a flexible machine learning (ML) approach that determines the cutoffs based on the data set qualities. However, the ML approach requires at least 20–30 similar data sets to run together so the patterns of errors can be learned. Here, we have used the hard filtering approach. The filters remove the variants with low confidence (QUAL parameter), the variants with low coverage (QD parameter), the variants with strand bias (FS and SOR parameters), the variants with reads that have low mapping quality (MQ and MQRankSum parameters— applied only to SNPs filtering) and the variants that are only detected at the end of the reads (ReadPosRankSum). (c) Merge SNPs and indels variants back together: java -jar picard.jar MergeVcfs \ I=vcf/final_snps_filtered.vcf.gz \ I=vcf/final_indels_filtered.vcf.gz \ O=vcf/filtered_comb.vcf.gz

16. Normalize the variants to make the representation of the variants consistent [27]: bcftools norm –m - -f ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ -O z vcf/filtered_comb.vcf.gz > final_report/filtered_cohort_norm.vcf.gz

This step splits multiallelic sites into multiple rows and leftalign and normalizes the indels. The VCF file generated in this step is the final filtered non-annotated *.vcf file that this workflow will produce. After this file is produced (filtered_cohort_norm.vcf.gz in this example), intermediate *.vcf files generated in the previous steps can be deleted. It is worth keeping an unannotated *.

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vcf file in the final_report folder in case the need for reannotation arises. 17. Annotate the variants using VEP. Before running this step, specify the full path to the location, where the VEP plugins and cache directory to be used are installed (the cache directory location can be chosen when installing VEP; by default, it is the same location as VEP): vep=/path/to/vep dir_cache=/path/to/vep_cache

Then, run the following command line: vep --cache --dir $dir \ --dir_cache $dir_cache \ --offline \ --fasta ref/GCA_000001405.15_GRCh38_no_alt_analysis_set.fna \ --species homo_sapiens \ --input_file final_report/filtered_cohort_norm.vcf.gz \ --output_file final_report/filtered_cohort_norm_annotated.vcf \ --format vcf \ --force_overwrite \ --vcf \ --no_check_variants_order \ --check_existing \ --freq_pop gnomAD \ --assembly GRCh38 \ --stats_file final_report/vep_stat.html \ --hgvs \ --variant_class \ --keep_csq \ --af_gnomad \ --polyphen p \ --sift p \ --symbol \ --total_length \ --max_af \ --plugin CADD,/path/to/whole_genome_SNVs.tsv.gz,/path/to/InDels.tsv.gz \ --plugin dbNSFP,/path/to/dbNSFP4.1a_grch38.gz,MutationTaster_pred \ --plugin ExACpLI,/path/to/ExACpLI_values.txt \ --plugin LoFtool \ --plugin DisGeNET,file=/path/to/all_variant_disease_pmid_associations_final.tsv.gz, \ disease=1 \ --plugin REVEL,/path/to/new_tabbed_revel_grch38.tsv.gz \ --plugin Mastermind,/path/to/mastermind_cited_variants_reference-2021.04. \ 02-grch38.vcf.gz,0,0,1 \ --fields "Gene,Feature,SYMBOL,Existing_variation,VARIANT_CLASS,Consequence, \ cDNA_position,CDS_position,Protein_position,Amino_acids,HGVSc,HGVSp,BIOTYPE, \

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IMPACT,CLIN_SIG,PolyPhen,SIFT,CADD_PHRED,CADD_RAW,MutationTaster_pred,REVEL, \ gnomAD_AF,MAX_AF,ExACpLI,LoFtool,DisGeNET_PMID,DisGeNET_SCORE,DisGeNET_disease, \ Mastermind_URL" \ --pick \ --pick_order rank,canonical,tsl \ --buffer_size 20000 \ --fork 4

Alongside the annotated vcf file, as an output, VEP will produce the statistics summary of the number of annotated variants and into which categories they fall. 18. Convert the vcf file to csv, add the custom Mitocarta annotation and apply initial filtering conditions. Python will be used to format an annotated vcf file and add the Mitocarta gene annotation. At this step we also automatically filter the variants based on the parameters that are unlikely to be associated with causative variants of mitochondrial diseases (Subheading 3.4 gives more details for these criteria). Remove variants that: • Have a population allele frequency of >0.05. • Are non-protein-altering consequences (intron variants, upstream and downstream gene variants, intergenic variants, 5′ and 3′ UTR variants; synonymous variants, non-coding transcript variants, pseudogene variants) (see Note 7). • Are not nuclear encoded mitochondrial genes according to Mitocarta (v3) (see Note 8). The initial filtering of the variants before prioritization is optional, but it simplifies the execution of the prioritization step, and typically reduces the number of variants from several million (based on the exome panel of 8 samples) to several thousand (Fig. 2) (see Note 9). The hash marks in the code indicate comments introduced into the python code: python3 import pandas as pd import numpy as np #indicate the location of the list with Mitocarta genes mitocarta3=’ref/Human.MitoCarta3.0.csv’ #Opening .vcf file as a dataframe in_df=pd.read_csv("final_report/filtered_cohort_norm_annotated.vcf", \ delimiter=’\t’, quotechar=’"’, quoting=2, comment=’#’, header=None) in_df.columns=header[0].split() df_A= in_df[[’#CHROM’,’POS’,’REF’,’ALT’,’INFO’,’FORMAT’]].copy() df_B=in_df[in_df.columns[9:]]

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1801 Pre-filtered annotated variants 290 Homozygous and heterozygous variants in Subject 1 94 protein altering variants

35 Rare variants (MAF≤0.01)

2 Homozygous variants

NAXD

Fig. 2 Funnel chart of prioritization of pre-filtered annotated variants in Subject 1 #Parsing the VEP annotation df_A[[’INFO’,’VEP’]]=in_df[’INFO’].str.split(’CSQ=’, expand=True) df_A[[’Gene’,’Feature’,’SYMBOL’,’Existing_variation’,’VARIANT_CLASS’, \ ’Consequence’,’cDNA_position’,’CDS_position’,’Protein_position’, \ ’Amino_acids’,’HGVSc’,’HGVSp’,’BIOTYPE’,’IMPACT’,’CLIN_SIG’,’PolyPhen’,’SIFT’, \ ’CADD_PHRED’,’CADD_RAW’,’MutationTaster_pred’,’REVEL’,’gnomAD_AF’,’MAX_AF’, \ ’ExACpLI’,’LoFtool’,’DisGeNET_PMID’,’DisGeNET_SCORE’,’DisGeNET_disease’, \ ’Mastermind_URL’]]=df_A[’VEP’].str.split(’|’,expand=True) #Making gnomAD frequencies numeric to later filter on them df_A[’gnomAD_AF’] = pd.to_numeric(df_A[’gnomAD_AF’]) #Adding MitoCarta annotation mitocarta=pd.read_csv(mitocarta3) df_A["In_MitoCarta3"]=df_A.Gene.isin(mitocarta.EnsemblGeneID)

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#Assigning Hom and Het to the variants in the samples def get_outcome(s): if ’0/1’ in s: return ’HET’ elif ’1/0’ in s: return ’HET’ elif ’1/2’ in s: return ’HET’ elif ’2/2’ in s: return ’HOM’ elif ’1/1’ in s: return ’HOM’ elif ’0/0’ in s: return ’REF’ else: return ’unknown’ flexcols = df_B.columns.tolist() new_cols = [] for col in flexcols: new_cols.append(in_df[col].apply(get_outcome).rename(col+’_ZYG’)) new_cols.append(in_df[col]) combined=pd.concat([df_A]+new_cols,axis=1) ##Final pivoting of the files: combined=combined.loc[ ~(combined[’gnomAD_AF’] >= 0.05)] combined=combined.loc[(combined[’In_MitoCarta3’] == True )] combined=combined.loc[(combined[’Consequence’].notna())] combined=combined.loc[~(combined[’Consequence’]==’intron_variant’)] combined=combined.loc[~(combined[’Consequence’]==’upstream_gene_variant’)] combined=combined.loc[~(combined[’Consequence’]==’downstream_gene_variant’)] combined=combined.loc[~(combined[’Consequence’]==’intergenic_variant’)] combined=combined.loc[~(combined[’Consequence’]==’5_prime_UTR_variant’)] combined=combined.loc[~(combined[’Consequence’]==’3_prime_UTR_variant’)] combined=combined.loc[~(combined[’Consequence’]==’synonymous_variant’)] combined=combined.loc[~(combined[’Consequence’]==’non_coding_transcript_variant’)] combined=combined.loc[~(combined[’Consequence’]==’intron_variant&non_coding_ \ transcript_variant’)] combined=combined.loc[~(combined[’Consequence’]==’non_coding_transcript_exon_ \ variant’)] combined.to_csv(’final_output/annotated.csv’, sep=’\t’) quit()

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Most of the steps described below can be performed automatically using the custom script; however, here, we describe a manual prioritization strategy that can be done in Microsoft Excel. The benefit of manual prioritization is that the analyst can review the variants and vary the stringency of prioritization based on the variants found. The example used is a panel of eight subjects from which we will describe filtering variants for Subject 1, who presented with a typical mitochondrial disease phenotype [23]. The additional seven subjects in the panel are treated as controls with unrelated phenotypes and were run through the same workflow (see Subheadings 1.2 and 3.2). Initial prefiltering, as described in step 18 of Subheading 3.2, has been applied to the samples, leaving 1801 variants for consideration. At each step in the filtering process, we are aiming to further reduce the number of variants (Fig. 2). 1. Convert the .csv file to .xls to enable all features: open the .csv output file in Microsoft Excel, in File go to Save as, File Format and select option Excel Workbook (.xlsx). 2. Consider apparent inheritance of the patient that you are analysing. In this example Subject 1’s family exhibits/fits with an autosomal recessive inheritance pattern [23]. We would, therefore, prioritize single homozygous variants or genes with two heterozygous variants (see Note 10). 3. Select HET and HOM changes in the patient of interest; for Subject 1 this is 290 variants (see Note 11). This is done because if running multiple samples together homozygous reference variants (REF) will be present in the sample at the positions, where control samples have variants. 4. Select protein-altering consequences of variants that would involve a change in amino acid: with VEP annotation this would include missense, stop loss, stop gain, frameshift. For a full list of the possible consequences defined by VEP see https://m.ensembl.org/info/genome/variation/prediction/ predicted_data.html . Consequences with Moderate and High impact should be considered first (see Note 12). After filtering Subjects 1’s data, 94 variants of interest remain. 5. Consider variants with a minor allele frequency of less than or equal to 1%, or not present in gnomAD (35 variants in Subject 1). Note if using gnomAD v2.1.1 when comparing to WES data (https://gnomad.broadinstitute.org/) as opposed to gnomAD v3.1.1: the v2 of this database contains a higher proportion of exomes, whereas v3.1.1 contains a higher proportion of genomes. If step 18 of Subheading 3.2 has been followed, then the minor allele frequencies according to gnomAD v2.1.1 are displayed in the column “gnomAD_AF” in the

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annotated *.csv file. MAX_AF from VEP output reports the highest allele frequency observed in any ethnic population from 1000 genomes, ESP or gnomAD. If any homozygotes are present in gnomAD, then exclude the variant on this basis. After this filter, only 33 variants are left for further investigation for Subject 1. 6. Use in silico prediction tools of pathogenicity such as Sorting Intolerant from Tolerant (SIFT; https://sift.bii.a-star.edu.sg/) [28], Polymorphism Phenotyping v2 (PolyPhen-2; http:// genetics.bwh.harvard.edu/pph2/) [29], Combined Annotation Dependent Depletion (CADD; https://cadd.bihealth. org/) [19] and REVEL [30] to prioritize potentially diseasecausing variants. These are displayed in the columns “SIFT”, “PolyPhen”, “CADD_PHRED” and “REVEL”, respectively, in the annotated csv file (see Note 13). SIFT takes into account protein conservation and classifies an amino acid change as tolerated or deleterious to protein function. PolyPhen-2 uses physical and evolutionary comparative considerations, and the amino acid changes are categorized as benign, possibly damaging, or probably damaging. CADD is a measure of variant deleteriousness built from 60+ genomic features; for CADD we consider a variant with a PHRED score greater than 25 as potentially pathogenic. REVEL is an ensemble method that aggregates the pathogenicity prediction of 13 separate tools and has been shown to have superior performance in variant pathogenicity prediction when compared to individual scoring systems [31]. Threshold scores for REVEL pathogenicity prediction are likely to be around ≥0.7 [21]. (Optional). Note that some genes are more tolerant to variants than others. The ExACoLI plugin reports the probability of a gene being loss-of-function intolerant [32]. LoFtool plugin also provides the quantification of genic intolerance to loss-of-function variants and the consequent susceptibility to disease, but reports the ratio of Loss-of-function (LoF) to synonymous mutations; the lower the LoFtool gene score percentile the more intolerant the gene is to functional variation [33]. ExACoLI and LoF values for the gene are reported in “ExACpLI” and “LoFtool” columns in the annotated csv file. 7. Check whether the canonical, high-confidence transcript is affected by the variant. The large number of alternatively spliced transcripts means that the same variant can lead to different consequences in each transcript encoded by the gene. In this situation, the transcript with the most severe consequences will be reported. As such, occasionally the bioinformatic pipeline will report the consequences of a change to the rare, non-canonical transcript

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instead of the high confident one. dbSNP, Ensembl and gnomAD v2.1.1 should be used to check whether the variant has similar consequences to the canonical, high confident transcript. Matched Annotation from NCBI and EMBL-EBI (MANE) provides one high-quality representative transcript per protein-coding gene that is well-supported by experimental data and represents the biology of the gene (see Note 14). 8. Investigate the function of the gene which is potentially affected by the prioritized variant. The gene function can be checked via Genecards (www.genecards.org) [34]; and any links to disease using Online Mendelian Inheritance in Man (OMIM; www.omim.org) [35]. Although OMIM is not exhaustive, its information can be supplemented by searching Pubmed (https://pubmed.ncbi.nlm.nih.gov/) for literature associated with the gene. Incorporating the plugins DisGeNET and MasterMind in the analysis output can also help with the task of collecting initial information about the gene/variant. MasterMind reports variants that have clinical evidence cited in the medical literature [36], while DisGeNET outputs the known genedisease associations [37]. The link to the MasterMind database of variants, the DisGeNET associated diseases and the confidence score of this disease association are reported in Mastermind_URL, DisGeNET_disease, and DisGeNET_SCORE columns of the annotated csv file, respectively. 9. Check whether the variant has been reported in ClinVar database [38]. ClinVar accumulates the submissions about clinically relevant variants that have been described previously in the patients and are assigned a status of pathogenic, benign, risk factor or insignificant. The” Clin_Sig” column in the annotated csv file reports whether the variant has been reported in ClinVar database [39]. However, it is important to check the ClinVar database itself for any associated phenotype assigned to the variant and any available evidence for the clinical interpretation. In our example of Subject 1, we used the filtering criteria and in silico pathogenicity prediction tools to narrow down the list of variants to two homozygous variants: rs369951364 (missense variant in NAXD gene) and rs28357685 (missense variant in mitochondrial CYB gene). However, the variant in the CYB gene, can be excluded, as this homozygous variant is listed as benign in the ClinVar database. 10. Present the bioinformatics and research evidence accumulated in steps 1–9 to the Clinical Diagnostic Team to produce the clinical report.

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For the case of Subject 1, used as an example in the chapter (Fig. 2), the report includes • the candidate gene (NAXD) variant annotation: cDNA (c.308C > T) and protein position (p.Pro103Leu) with the relevant RefSeq (NM_001242882. 1), • details on MAF from gnomAD (0.0000478), • in silico prediction results including PolyPhen (probably damaging), SIFT (deleterious) and CADD PHRED score (26.9). 3.4 Sanger Confirmation and Clinical Reporting

1. Check the read depth of the variant using the metrics from the GATK output. GATK reports the genotype characteristics of the variants in the following format: GT:AD:DP:GQ:PL. GT (genotype) is genotype in the allelic form; AD (allele depth) is the number of reads that support each of the reported alleles; DP (depth of coverage) is the number of filtered reads that support each of the reported alleles; GQ (genotype quality) is the Phred-scaled confidence that the genotype assignment (GT) is correct, it is capped at 99; PL (Phred-scaled likelihood) is “normalized” Phred-scaled likelihoods of the possible genotypes. For the NAXD variant in our example of Subject 1, the characteristics are 1/1:0,60:60:99:1928,178,0, indicating that it is a homozygous variant (1/1) which is covered by a sufficient number of reads (60); none of the reads has REF nucleotide in them (0,60); the genotype quality for this variant is extremely high (99). 2. (Optional) Visualize the variant in the genome browser, IGV. To visualize the variant in the genome browser the bam file covering the variant is required. It helps to make a separate bam file for the region of interest, instead of loading the bam file for the whole sample. Here, we extract the region around the NAXD gene (chr13:110,613,460-110,641,996) from the Subject 1 bam file:

samtools view -b bam/subject1_dedup.bam ‘chr13:110,613,460-110,641,996’ > \ final_output/NAXD_subject1.bam samtools index final_output/NAXD_subject1.bam

Both bam file and its index (file extension .bam.bai) are required for loading in IGV. 3. Where no robust series of checks or end-to-end robotics are in place to abrogate the possibility of sample mixup, Sanger sequencing should be undertaken to validate the presence of the variant prior to reporting.

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Notes 1. Due to the lack of patient consent, it is not possible to make the discussed data set from subject 1 publicly available. However, this data set was selected as a representative example of the sequencing information routinely obtained in our laboratory and prioritization strategies to identify the disease-causing variant. 2. Since the exome capture requires an enrichment step, the results of the enrichment will depend on the quality of DNA and the exome capture kit in use. However, exome enrichment is usually performed by the facility providing the highthroughput sequencing services and there might not be a choice of what kit is being used. As such, extra care should be taken to ensure that the quality of genomic DNA extracted from the patient sample material is high. 3. If the cost of sequencing is a constraint or tissue and DNA resampling for repeat sequencing is an issue, in most cases, it is possible to archive the adequate results with lower than recommended 100× coverage of whole-exome sequencing. Our laboratory routinely obtains 60–120× median coverage, and the data set discussed in this chapter has 49× median coverage, with 59.4 mln sequenced paired reads obtained. Recent publications suggest that even 30× coverage may be enough [40] when using a sufficiently advanced variant caller. 4. Sequencing facilities provide the sequenced data in the form of fastq files. As a rule, fastq files are output in compressed form (files ending with fastq.gz extension) and with the technical sequences used for making libraries (adapters and barcodes) already removed. In its compressed form the sequencing file for a sample is usually several Gb in size. 5. Often, sequence libraries are split across several lanes of a flow cell, producing more than one fastq file per sample for single end sequencing or >2 fastq files for paired-end sequencing. These files need to be concatenated using the command line. In the case of paired end sequencing, the order of fastq file concatenation must be the same for the forward and reverse reads. 6. If the quality of the sequenced files deviates from the expected, the most appropriate course of action is to repeat the sequencing. However, when such an option is not available, the sequenced reads can be trimmed by Trimmomatic [41] or similar tools to remove the adapters and low-quality bases from the start and ends of the reads.

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7. Though at this stage we exclude the variants that do not alter proteins, it is possible that these variants can still be causative variants for mitochondrial diseases [42]. In general, confirmation that a non-protein altering variant is a causative variant requires non-routine work and extensive experimental validation. 8. Note that it is possible for the patient to exhibit a mitochondrial-like disease phenotype, while the causative gene does not belong to the list of MitoCarta genes [43]. 9. These commands are written in Python; the first line of the code, python3, invokes the python environment. Note that in Python the syntax blocks (e.g., if statements, for loops, functions etc) are delimited by indentation, so extra attention should be paid to have consistent whitespaces when copying/ typing the code. 10. For consanguineous families consider homozygous variants first. 11. The variants present will receive HET, HOM or REF annotation if the final vcf file (filtered_cohort_norm_annotated.vcf) has been subjected to the formatting in step 18 of Subheading 3.2. Otherwise, the variant genotype will be encoded as allele values (0/0 for REF, 0/1 for HET and so on according to vcf format specification) and filtering should be adjusted accordingly. 12. Note that the majority of the variants that are not proteinaltering have been excluded at step 18 of Subheading 3.2. Additional manual filtering of consequences allows the user to vary the stringency of the variant consequences. 13. Please note that sometimes the prediction tools will give different pathogenicity predictions for the same variant, these cases should be inspected manually on a case-by-case basis. Predictions for SNPs are overall in a more mature state than the ones for indels. 14. The instructions of how to access MANE transcript annotation, depending on the genome browser in use, are available at https://www.ncbi.nlm.nih.gov/refseq/MANE/.

Acknowledgments Work in our laboratories is supported by the Wellcome Centre for Mitochondrial Research (203105/Z/16/Z), the Medical Research Council (MRC) International Centre for Genomic Medicine in Neuromuscular Disease (MR/S005021/1), the National Institute of Health Research (NIHR) Biomedical Research Centre

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in Age and Age Related Diseases award to the Newcastle upon Tyne Hospitals National Health Service (NHS) Foundation, the Lily Foundation, the Pathology Society and the NHS Highly Specialised Service for Rare Mitochondrial Disorders. CLA is supported by the National Institute for Health Research (NIHR) PostDoctoral Fellowship (PDF-2018-11-ST2-021). Figure 1 was created with BioRender.com. The views expressed in this publication are those of the author(s) and not necessarily those of the NHS, the NIHR, or the Department of Health and Social Care. References 1. Stenton SL, Prokisch H (2018) Advancing genomic approaches to the molecular diagnosis of mitochondrial disease. Essays Biochem 62(3):399–408 2. Stenton SL, Shimura M, PiekutowskaAbramczuk D, Freisinger P, Distelmaier F, Mayr JA et al (2021) Diagnosing pediatric mitochondrial disease: lessons from 2,000 exomes. medRxiv 3. Alston CL, Stenton SL, Hudson G, Prokisch H, Taylor RW (2021) The genetics of mitochondrial disease: dissecting mitochondrial pathology using multi-omic pipelines. J Pathol 254(4):430–442 4. Ye´pez VA, Gusic M, Kopajtich R, Mertes C, Smith NH, Alston CL et al (2021) Clinical implementation of RNA sequencing for Mendelian disease diagnostics. medRxiv 5. Kopajtich R, Smirnov D, Stenton SL, Loipfinger S, Meng C, Scheller IF et al (2021) Integration of proteomics with genomics and transcriptomics increases the diagnostic rate of Mendelian disorders. medRxiv 6. Thompson K, Collier JJ, Glasgow RIC, Robertson FM, Pyle A, Blakely EL et al (2020) Recent advances in understanding the molecular genetic basis of mitochondrial disease. J Inherit Metab Dis 43(1):36–50 7. Schwarze K, Buchanan J, Taylor JC, Wordsworth S (2018) Are whole-exome and wholegenome sequencing approaches cost-effective? A systematic review of the literature. Genet Med 20(10):1122–1130 8. Fang H, Wu Y, Narzisi G, O’Rawe JA, Barron LT, Rosenbaum J et al (2014) Reducing INDEL calling errors in whole genome and exome sequencing data. Genome Med 6(10): 89 9. Holt IJ, Harding AE, Morgan-Hughes JA (1988) Deletions of muscle mitochondrial

DNA in patients with mitochondrial myopathies. Nature 331(6158):717–719 10. Grady JP, Pickett SJ, Ng YS, Alston CL, Blakely EL, Hardy SA et al (2018) mtDNA heteroplasmy level and copy number indicate disease burden in m.3243A>G mitochondrial disease. EMBO Mol Med 10(6) 11. Hamzeh AR, Andrews TD, Field MA (2021) Detecting causal variants in Mendelian disorders using whole-genome sequencing. Methods Mol Biol 2243:1–25 12. Poplin R, Ruano-Rubio V, DePristo MA, Fennell TJ, Carneiro MO, Van der Auwera GA et al (2017) Scaling accurate genetic variant discovery to tens of thousands of samples. bioRxiv 13. Rimmer A, Phan H, Mathieson I, Iqbal Z, Twigg SRF, Consortium WGS et al (2014) Integrating mapping-, assemblyand haplotype-based approaches for calling variants in clinical sequencing applications. Nat Genet 46(8):912–918 14. Afgan E, Baker D, Batut B, van den Beek M, Bouvier D, Cech M et al (2018) The Galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2018 update. Nucleic Acids Res 46(W1):W537–WW44 15. Wang K, Li M, Hakonarson H (2010) ANNOVAR: functional annotation of genetic variants from high-throughput sequencing data. Nucleic Acids Res 38(16):e164 16. McLaren W, Gil L, Hunt SE, Riat HS, Ritchie GR, Thormann A et al (2016) The ensembl variant effect predictor. Genome Biol 17(1): 122 17. Cingolani P, Platts A, Wang le L, Coon M, Nguyen T, Wang L et al (2012) A program for annotating and predicting the effects of single nucleotide polymorphisms, SnpEff: SNPs in the genome of Drosophila melanogaster strain w1118; iso-2; iso-3. Fly (Austin) 6(2):80–92

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30. Ioannidis NM, Rothstein JH, Pejaver V, Middha S, McDonnell SK, Baheti S et al (2016) REVEL: an ensemble method for predicting the pathogenicity of rare missense variants. Am J Hum Genet 99(4):877–885 31. Tian Y, Pesaran T, Chamberlin A, Fenwick RB, Li S, Gau CL et al (2019) REVEL and BayesDel outperform other in silico meta-predictors for clinical variant classification. Sci Rep 9(1): 12752 32. Lek M, Karczewski KJ, Minikel EV, Samocha KE, Banks E, Fennell T et al (2016) Analysis of protein-coding genetic variation in 60,706 humans. Nature 536(7616):285–291 33. Fadista J, Oskolkov N, Hansson O, Groop L (2017) LoFtool: a gene intolerance score based on loss-of-function variants in 60 706 individuals. Bioinformatics 33(4):471–474 34. Stelzer G, Rosen N, Plaschkes I, Zimmerman S, Twik M, Fishilevich S et al (2016) The GeneCards suite: from gene data mining to disease genome sequence analyses. Curr Protoc Bioinformatics 54:1 30 1–1 3 35. Amberger JS, Bocchini CA, Scott AF, Hamosh A (2019) OMIM.org: leveraging knowledge across phenotype-gene relationships. Nucleic Acids Res 47(D1):D1038–D1D43 36. Chunn LM, Nefcy DC, Scouten RW, Tarpey RP, Chauhan G, Lim MS et al (2020) Mastermind: a comprehensive genomic association search engine for empirical evidence curation and genetic variant interpretation. Front Genet 11:577152 37. Pinero J, Ramirez-Anguita JM, Sauch-Pitarch J, Ronzano F, Centeno E, Sanz F et al (2020) The DisGeNET knowledge platform for disease genomics: 2019 update. Nucleic Acids Res 48(D1):D845–DD55 38. Landrum MJ, Lee JM, Benson M, Brown G, Chao C, Chitipiralla S et al (2016) ClinVar: public archive of interpretations of clinically relevant variants. Nucleic Acids Res 44(D1): D862–D868 39. Harrison SM, Riggs ER, Maglott DR, Lee JM, Azzariti DR, Niehaus A et al (2016) Using ClinVar as a resource to support variant interpretation. Curr Protoc Hum Genet 89: 8 16 1–8 8 23 40. Pei S, Liu T, Ren X, Li W, Chen C, Xie Z (2021) Benchmarking variant callers in nextgeneration and third-generation sequencing analysis. Brief Bioinform 22(3):bbaa148 41. Bolger AM, Lohse M, Usadel B (2014) Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30(15): 2114–2120

Genomic Strategies in Mitochondrial Diagnostics 42. Zeng Z, Bromberg Y (2019) Predicting functional effects of synonymous variants: a systematic review and perspectives. Front Genet 10: 914 43. Pyle A, Nightingale HJ, Griffin H, Abicht A, Kirschner J, Baric I et al (2015) Respiratory chain deficiency in nonmitochondrial disease. Neurol Genet 1(1):e6 44. Kemp JP, Smith PM, Pyle A, Neeve VC, Tuppen HA, Schara U et al (2011) Nuclear factors involved in mitochondrial translation cause a subgroup of combined respiratory chain deficiency. Brain 134(Pt 1):183–195 45. Lieber DS, Calvo SE, Shanahan K, Slate NG, Liu S, Hershman SG et al (2013) Targeted

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exome sequencing of suspected mitochondrial disorders. Neurology 80(19):1762–1770 46. Taylor RW, Pyle A, Griffin H, Blakely EL, Duff J, He L et al (2014) Use of wholeexome sequencing to determine the genetic basis of multiple mitochondrial respiratory chain complex deficiencies. JAMA 312(1): 68–77 47. Hengel H, Hannan SB, Dyack S, MacKay SB, Schatz U, Fleger M et al (2021) Bi-allelic lossof-function variants in BCAS3 cause a syndromic neurodevelopmental disorder. Am J Hum Genet 108(6):1069–1082

Chapter 28 Mitochondrial DNA Enrichment for Sensitive Next-Generation Sequencing Shilan Wu, Matthew J. Longley, Scott A. Lujan, Thomas A. Kunkel, and William C. Copeland Abstract Mitochondrial DNA (mtDNA) encodes components essential for cellular respiration. Low levels of point mutations and deletions accumulate in mtDNA during normal aging. However, improper maintenance of mtDNA results in mitochondrial diseases, stemming from progressive loss of mitochondrial function through the accelerated formation of deletions and mutations in mtDNA. To better understand the molecular mechanisms underlying the creation and propagation of mtDNA deletions, we developed the LostArc next-generation DNA sequencing pipeline to detect and quantify rare mtDNA species in small tissue samples. LostArc procedures are designed to minimize PCR amplification of mtDNA and instead achieve enrichment of mtDNA by selective destruction of nuclear DNA. This approach leads to costeffective, high-depth sequencing of mtDNA with a sensitivity sufficient to identify one mtDNA deletion per million mtDNA circles. Here, we describe detailed protocols for isolation of genomic DNA from mouse tissues, enrichment of mtDNA through enzymatic destruction of linear nuclear DNA, and preparation of libraries for unbiased next-generation sequencing of mtDNA. Key words Mitochondrial DNA, DNA deletions, Next-Generation Sequencing, Mitochondrial disease, POLG, Mitochondrial DNA Replication

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Introduction Human mitochondria possess a circular 16,569 bp doublestranded DNA genome that supports cellular respiration by encoding 37 genes essential for electron transport and oxidative phosphorylation. Mitochondrial dysfunction can result from depletion of mitochondrial DNA (mtDNA), as well as through the formation of point mutations, deletions, and duplications in mtDNA. Depletion of mtDNA is linked to early onset, often fatal mitochondrial diseases [1, 2]. Maternally inherited defects in human mtDNA were first reported in 1988 as point mutations associated with Leber’s hereditary optic neuropathy [3] and as mtDNA deletions up to 7 kb in length in mitochondrial myopathies [4]. Since that time,

Thomas J. Nicholls et al. (eds.), Mitochondrial DNA: Methods and Protocols, Methods in Molecular Biology, vol. 2615, https://doi.org/10.1007/978-1-0716-2922-2_28, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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nearly 700 point mutations and several hundred deletions in mtDNA have been associated with various forms of mitochondrial disease. Eukaryotic cells can contain thousands of copies of mtDNA, and both wild-type and mutated mtDNA sequences coexist as a heteroplasmic mixture in the same cell. Low levels of mtDNA heteroplasmy have been detected in healthy individuals [5]. Interestingly, both point mutations [6] and deletions [7, 8] were found to accumulate in mtDNA as a function of aging, and such changes may participate in the normal aging process. Progressive external ophthalmoplegia (PEO) is a familial condition characterized by weakness or paralysis of the external eye muscles, ptosis, and progressive skeletal muscle weakness. In 1989, multiple mtDNA deletions were first identified in patients with PEO and were implicated in impaired mitochondrial function [9]. Autosomal inheritance of the first PEO disease allele was linked to chromosomal locus 10q23.3–24.3 [10], which was later identified as the TWNK gene encoding Twinkle helicase [11]. Soon thereafter, autosomal PEO loci for ANT1 encoding the adenine nucleotide translocator [12] and POLG encoding the catalytic subunit of the mitochondrial DNA polymerase γ [13] were identified. Mutations in nuclear genes encoding components of the mitochondrial replisome co-segregate with mitochondrial diseases [14], indicating that faulty mtDNA replication can cause a variety of heritable mitochondrial diseases [15]. The POLG gene product possesses both DNA polymerase and 3′–5′ exonuclease functions and combines with the POLG2-encoded processivity accessory factor to form the DNA polymerase γ holoenzyme. The Pol γ holoenzyme is the sole replicative DNA polymerase responsible for duplicating the mitochondrial genome [16, 17]. Pol γ works in concert with the Twinkle helicase, SSBP1, and other accessory factors [15, 18] to carry out efficient mtDNA replication. Mutations in POLG represent the most common source of inherited mitochondrial diseases, and more than 300 pathogenic mutations of POLG have been reported (https://tools.niehs.nih.gov/polg/). POLG mutations cause a spectrum of disorders with onset ranging from early childhood in myocerebrohepatopathy spectrum disorders or Alpers–Huttenlocher syndrome to adult presentation of ataxia–neuropathy spectrum or PEO [19, 20]. While early onset POLG disorders stem from depletion of mtDNA, later onset POLG disorders frequently feature the accumulation of mtDNA deletions that compromise mitochondrial function [20]. To gain insight into the molecular mechanisms underlying formation of mtDNA deletions, we developed LostArc, a highly sensitive pipeline to delineate the position, length, sequence context, and abundance of mtDNA deletions in human tissue samples [21]. MtDNA represents