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Methods in Molecular Biology 2276

Volkmar Weissig Marvin Edeas Editors

Mitochondrial Medicine Volume 2: Assessing Mitochondria Second Edition

METHODS IN MOLECULAR BIOLOGY

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

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

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

Mitochondrial Medicine Volume 2: Assessing Mitochondria Second Edition

Edited by

Volkmar Weissig Department of Pharmaceutical Sciences, Midwestern University, Glendale, AZ, USA

Marvin Edeas Cochin Hospital, Cochin Institute, INSERM U1016, PARIS, France

Editors Volkmar Weissig Department of Pharmaceutical Sciences Midwestern University Glendale, AZ, USA

Marvin Edeas Cochin Hospital Cochin Institute, INSERM U1016 PARIS, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1265-1 ISBN 978-1-0716-1266-8 (eBook) https://doi.org/10.1007/978-1-0716-1266-8 © Springer Science+Business Media, LLC, part of Springer Nature 2021 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 Springer 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 It is our distinct pleasure to present the second edition of MiMB Mitochondrial Medicine to the ever increasing number of scientists and physicians who are as fascinated by this tiny organelle as we are. We started working on the first edition in September 2014 and were able to bring about one year later two volumes with a total of 70 chapters to the market. As of today (July 2020), 195K downloads have been recorded for Volume I1 and 90K downloads for volume II2. In light of the rapidly growing and expanding field of Mitochondrial Medicine, we readily accepted the invitation to compile a second edition, which we started to work on in March 2019. This second edition as offered here involves a total of 88 chapters with 45 of them written by new contributors who were not part of our first edition. The first and second editions combined subsequently present work from 115 mitochondrial laboratories from around the globe. We therefore believe these five volumes combined to be the most comprehensive source of know-how in the wide-ranging field of Mitochondrial Medicine. Dividing 87 chapters equally over three volumes proved to be a bit challenging. We chose the subtitle Targeting Mitochondria for volume I, Assessing Mitochondria for volume II, and Manipulating Mitochondria and Disease Specific Approaches for volume III while of course being well aware of significant overlaps between these three areas of research. For example, it is quite obvious that mitochondria are being targeted for the purpose of either assessing them or to manipulate them. We therefore ask all authors not to be too critical regarding the placement of their particular chapter. The reader we believe will anyway choose to download a chapter of his/her interest quite independently of its placement in one of the three volumes. All chapters in these three volumes were written for graduate students, postdoctoral associates, independent investigators in academia and industry as well as physicians by leading experts in their particular field. We are extremely grateful to them for having found the time to either update their chapter from the first edition or to write a new chapter. We will not forget that for many if not all of our contributors the worldwide COVID-19 pandemic posed additional and unexpected hurdles towards finishing their manuscript in due time. Thank you to all! The idea for our original book proposal leading to the first edition of MiMB Mitochondrial Medicine originated in our efforts to organize a series of annual conferences on Targeting Mitochondria (www.targeting-mitochondria.com), the tenth one of which meanwhile has taken place in November 2019 in Berlin, Germany. Due to the ongoing pandemic, our 11th conference (October 29–30, 2020) will be a virtual one but we are sure it will not be less exciting than all the previous editions. Last but not least we would like to sincerely thank John Walker, the series editor of Methods in Molecular Biology, for having invited us to compile this second edition and for his

1 2

https://link.springer.com/book/10.1007/978-1-4939-2257-4. https://link.springer.com/book/10.1007/978-1-4939-2288-8.

Preface

unlimited guidance and help throughout the entire process. We also owe sincere thanks to Patrick Marton, the Executive Editor of the Springer Protocol Series, for always having been available in assisting us throughout the entire project. Glendale, AZ, USA Paris, France

Volkmar Weissig Marvin Edeas

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

1 Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations, Pitfalls, and Tomorrow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Naig Gueguen, Guy Lenaers, Pascal Reynier, Volkmar Weissig, and Marvin Edeas 2 Preparation of “Functional” Mitochondria: A Challenging Business. . . . . . . . . . . Stefan Lehr, Sonja Hartwig, and Jorg Kotzka 3 Isolation and Quality Control of Functional Mitochondria . . . . . . . . . . . . . . . . . . . Sonja Hartwig, Jorg Kotzka, and Stefan Lehr 4 Purification of Functional Platelet Mitochondria Using a Discontinuous Percoll Gradient . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jacob L. Léger, Nicolas Pichaud, and Luc H. Boudreau 5 Mechanical Permeabilization as a New Method for Assessment of Mitochondrial Function in Insect Tissues. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alessandro Gaviraghi, Yan Aveiro, Stephanie S. Carvalho, Rodiesley S. Rosa, Matheus P. Oliveira, and Marcus F. Oliveira 6 Analysis of Mitochondrial Retrograde Signaling in Yeast Model Systems . . . . . . . Nicoletta Guaragnella, Maša Ždralević, Zdena Palková, and Sergio Giannattasio 7 Native Gel Electrophoresis and Immunoblotting to Analyze Electron Transport Chain Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gisela Beutner and George A. Porter Jr. 8 Measuring Mitochondrial Hydrogen Peroxide Levels and Glutathione Redox Equilibrium in Drosophila Neuron Subtypes Using Redox-Sensitive Fluorophores and 3D Imaging. . . . . . . . . . . . . . . . . . . . . . . Lori M. Buhlman, Petros P. Keoseyan, Kathryn Houlihan, and Amber N. Juba 9 Assessment of Mitochondrial Cell Metabolism by Respiratory Chain Electron Flow Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Flavia Radogna, Déborah Gérard, Mario Dicato, and Marc Diederich 10 Whole-Cell and Mitochondrial dNTP Pool Quantification from Cells and Tissues. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Juan C. Landoni, Liya Wang, and Anu Suomalainen 11 Single-Particle Tracking Method in Fluorescence Microscopy to Monitor Bioenergetic Responses of Individual Mitochondria . . . . . . . . . . . . . . Camille Colin, Emmanuel Suraniti, Emma Abell, Audrey Sémont, Neso Sojic, Philippe Diolez, and Stéphane Arbault

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Contents

Investigation of Mitochondrial ADP-Ribosylation Via Immunofluorescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ann-Katrin Hopp and Michael O. Hottiger Assessment of Mitochondrial Ca2+ Uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . András T. Deak, Claire Jean-Quartier, Alexander I. Bondarenko, Lukas N. Groschner, Roland Malli, Wolfgang F. Graier, and Markus Waldeck-Weiermair Assessment of Mitochondrial Membrane Potential and NADH Redox State in Acute Brain Slices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrey Y. Vinokurov, Viktor V. Dremin, Gennadii A. Piavchenko, Olga A. Stelmashchuk, Plamena R. Angelova, and Andrey Y. Abramov Evaluation of Mitochondria Content and Function in Live Cells by Multicolor Flow Cytometric Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hsiu-Han Fan, Tsung-Lin Tsai, Ivan L. Dzhagalov, and Chia-Lin Hsu Analysis of Mitochondrial Dysfunction During Cell Death . . . . . . . . . . . . . . . . . . . Vladimir Gogvadze and Boris Zhivotovsky Modified Blue Native Gel Approach for Analysis of Respiratory Supercomplexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sergiy M. Nadtochiy, Megan Ngai, and Paul S. Brookes Patch-Clamp Recording of the Activity of Ion Channels in the Inner Mitochondrial Membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Piotr Bednarczyk, Rafał P. Kampa, Shur Gałecka, Aleksandra Se˛k, Agnieszka Walewska, and Piotr Koprowski Assessment of Mitochondrial Protein Glutathionylation as Signaling for CO Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ana S. Almeida, Cláudia Figueiredo-Pereira, and Helena L. A. Vieira 3D Optical Cryo-Imaging Method: A Novel Approach to Quantify Renal Mitochondrial Bioenergetics Dysfunction . . . . . . . . . . . . . . . . . Shima Mehrvar, Amadou K. S. Camara, and Mahsa Ranji Simultaneous Quantification of Mitochondrial ATP and ROS Production Using ATP Energy Clamp Methodology . . . . . . . . . . . . . . . . . . . . . . . . Liping Yu, Brian D. Fink, and William I. Sivitz High-Throughput Image Analysis of Lipid-Droplet-Bound Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nathanael Miller, Dane Wolf, Nour Alsabeeh, Kiana Mahdaviani, Mayuko Segawa, Marc Liesa, and Orian S. Shirihai Cell Energy Budget Platform for Multiparametric Assessment of Cell and Tissue Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dmitri B. Papkovsky and Alexander V. Zhdanov Fluorescence-Based Assay For Measuring OMA1 Activity. . . . . . . . . . . . . . . . . . . . Julia Tobacyk and Lee Ann MacMillan-Crow Studying Mitochondrial Network Formation by In Vivo and In Vitro Reconstitution Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wanqing Du, Xiangjun Di, and Qian Peter Su

165 173

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Contents

Extraction of Functional Mitochondria Based on Membrane Stiffness . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Md Habibur Rahman, Qinru Xiao, Shirui Zhao, An-Chi Wei, and Yi-Ping Ho A Protocol for Untargeted Metabolomic Analysis: From Sample Preparation to Data Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amanda L. Souza and Gary J. Patti A Method for Analysis of Nitrotyrosine-Containing Proteins by Immunoblotting Coupled with Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . Matej Kohutiar and Adam Eckhardt In Vivo Visualization and Quantification of Mitochondrial Morphology in C. elegans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . R. de Boer, R. L. Smith, W. H. De Vos, E. M. M. Manders, and H. van der Spek Assessing Impact of Platinum Complexes on Mitochondrial Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Suxing Jin and Xiaoyong Wang In Silico Modeling of the Mitochondrial Pumping Complexes with Markov State Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roger Springett Monitoring the Mitochondrial Presequence Import Pathway In Living Mammalian Cells with a New Molecular Biosensor. . . . . . . . . . . . . . . . . Maxime Jacoupy, Emeline Hamon-Keromen, and Olga Corti

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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441 453

Contributors EMMA ABELL • Univ. Bordeaux, INSERM, Centre de recherche Cardio- Thoracique de Bordeaux, U1045 & IHU Liryc, Electrophysiology and Heart Modeling Institute, Fondation Bordeaux Université, Bordeaux, France; CHU de Bordeaux, Bordeaux, France ANDREY Y. ABRAMOV • Orel State University, Orel, Russia; UCL Queen Square Institute of Neurology, London, UK ANA S. ALMEIDA • CEDOC, Faculdade de Ciência Médicas/NOVA Medical School, Universidade Nova de Lisboa, Lisboa, Portugal NOUR ALSABEEH • Department of Physiology, Faculty of Medicine, Kuwait University, Kuwait City, Kuwait PLAMENA R. ANGELOVA • UCL Queen Square Institute of Neurology, London, UK STÉPHANE ARBAULT • NSysA group, Univ. Bordeaux, CNRS, INP Bordeaux, ISM, UMR 5255, Talence, France YAN AVEIRO • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil PIOTR BEDNARCZYK • Department of Physics and Biophysics, Institute of Biology, Warsaw University of Life Sciences—SGGW, Warsaw, Poland GISELA BEUTNER • Department of Pediatrics-Division Cardiology, University of Rochester, Rochester, NY, USA ALEXANDER I. BONDARENKO • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria LUC H. BOUDREAU • Department of Chemistry and Biochemistry, Universite de Moncton, Moncton, NB, Canada PAUL S. BROOKES • Department of Anesthesiology, University of Rochester Medical Center, Rochester, NY, USA LORI M. BUHLMAN • Arizona College of Graduate Studies, Midwestern University, Glendale, AZ, USA AMADOU K. S. CAMARA • Department of Anesthesiology and Anesthesia Research, Medical College of Wisconsin, Wauwatosa, WI, USA STEPHANIE S. CARVALHO • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil CAMILLE COLIN • NSysA group, Univ. Bordeaux, CNRS, INP Bordeaux, ISM, UMR 5255, Talence, France; Univ. Bordeaux, INSERM, Centre de recherche Cardio- Thoracique de Bordeaux, U1045 & IHU Liryc, Electrophysiology and Heart Modeling Institute, Fondation Bordeaux Université, Bordeaux, France; CHU de Bordeaux, Bordeaux, France OLGA CORTI • Sorbonne Université, Institut du Cerveau (ICM), Inserm U1127, CNRS UMR 7225, Paris, France R. DE BOER • Molecular Biology & Microbial Food Safety, Swammerdam Institute for Life Sciences (SILS), Faculty of Science (FNWI), University of Amsterdam, Amsterdam, The Netherlands W. H. DE VOS • Laboratory of Cell Biology and Histology, Department of Veterinary Sciences, Antwerp University, Antwerp, Belgium; Cell Systems and Imaging Research Group, Department of Molecular Biotechnology, Ghent University, Ghent, Belgium

xii

Contributors

ANDRÁS T. DEAK • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria XIANGJUN DI • Institute for Biomedical Materials & Devices (IBMD), Faculty of Science, University of Technology Sydney, Sydney, NSW, Australia MARIO DICATO • Laboratoire de Biologie Moléculaire et Cellulaire du Cancer, Hôpital Kirchberg, Luxembourg, Luxembourg MARC DIEDERICH • College of Pharmacy, Seoul National University, Seoul, South Korea PHILIPPE DIOLEZ • Univ. Bordeaux, INSERM, Centre de recherche Cardio- Thoracique de Bordeaux, U1045 & IHU Liryc, Electrophysiology and Heart Modeling Institute, Fondation Bordeaux Université, Bordeaux, France; CHU de Bordeaux, Bordeaux, France VIKTOR V. DREMIN • Orel State University, Orel, Russia; Aston University, Birmingham, UK WANQING DU • State Key Laboratory of Membrane Biology, Tsinghua University-Peking University Joint Center for Life Sciences, School of Life Sciences, Tsinghua University, Beijing, China IVAN L. DZHAGALOV • Institute of Microbiology and Immunology, National Yang-Ming University, Taipei, Taiwan ADAM ECKHARDT • Department of Translational Metabolism, Institute of Physiology, Academy of Sciences of the Czech Republic, Prague, Czech Republic MARVIN EDEAS • Université de Paris, INSERM U1016, Institut Cochin, CNRS UMR8104, Paris, France; Laboratory of Excellence GR-Ex, Paris, France HSIU-HAN FAN • Institute of Microbiology and Immunology, National Yang-Ming University, Taipei, Taiwan CLÁUDIA FIGUEIREDO-PEREIRA • CEDOC, Faculdade de Ciência Médicas/NOVA Medical School, Universidade Nova de Lisboa, Lisboa, Portugal BRIAN D. FINK • Division of Endocrinology and Metabolism, Department of Internal Medicine, The University of Iowa, Iowa City, IA, USA SHUR GAŁECKA • Laboratory of Intracellular Ion Channels, Nencki Institute of Experimental Biology, Warsaw, Poland ALESSANDRO GAVIRAGHI • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil DÉBORAH GÉRARD • Laboratoire de Biologie Moléculaire et Cellulaire du Cancer, Hôpital Kirchberg, Luxembourg, Luxembourg SERGIO GIANNATTASIO • Institute of Biomembranes, Bioenergetics and Molecular Biotechnologies, CNR, Bari, Italy VLADIMIR GOGVADZE • Division of Toxicology, Institute of Environmental Medicine, Karolinska Institutet, Stockholm, Sweden; Faculty of Medicine, MV Lomonosov Moscow State University, Moscow, Russia WOLFGANG F. GRAIER • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria LUKAS N. GROSCHNER • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria NICOLETTA GUARAGNELLA • Institute of Biomembranes, Bioenergetics and Molecular Biotechnologies, CNR, Bari, Italy; Department of Biosciences, Biotechnology and Biopharmaceutics, University of Bari “A. Moro”, Bari, Italy

Contributors

xiii

NAIG GUEGUEN • UMR CNRS 6015-INSERM U1083, MitoVasc Institute, University of Angers, Angers, France; Department of Biochemistry and Genetics, University Hospital of Angers, Angers, France EMELINE HAMON-KEROMEN • Sorbonne Université, Institut du Cerveau (ICM), Inserm U1127, CNRS UMR 7225, Paris, France SONJA HARTWIG • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center at the Heinrich-Heine-University Duesseldorf, Leibniz Center for Diabetes Research, Duesseldorf, Germany; German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany YI-PING HO • Department of Biomedical Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China; Centre for Novel Biomaterials, The Chinese University of Hong Kong, Hong Kong SAR, China; Shun Hing Institute of Advanced Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China; The Ministry of Education Key Laboratory of Regeneration Medicine, The Chinese University of Hong Kong, Hong Kong SAR, China ANN-KATRIN HOPP • Department of Molecular Mechanisms of Disease (DMMD), University of Zurich, Zurich, Switzerland MICHAEL O. HOTTIGER • Department of Molecular Mechanisms of Disease (DMMD), University of Zurich, Zurich, Switzerland KATHRYN HOULIHAN • Arizona College of Graduate Studies, Midwestern University, Glendale, AZ, USA CHIA-LIN HSU • Institute of Microbiology and Immunology, National Yang-Ming University, Taipei, Taiwan MAXIME JACOUPY • Sorbonne Université, Institut du Cerveau (ICM), Inserm U1127, CNRS UMR 7225, Paris, France CLAIRE JEAN-QUARTIER • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria SUXING JIN • State Key Laboratory of Coordination Chemistry, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing, P. R. China AMBER N. JUBA • Arizona College of Graduate Studies, Midwestern University, Glendale, AZ, USA RAFAŁ P. KAMPA • Department of Physics and Biophysics, Institute of Biology, Warsaw University of Life Sciences—SGGW, Warsaw, Poland; Laboratory of Intracellular Ion Channels, Nencki Institute of Experimental Biology, Warsaw, Poland PETROS P. KEOSEYAN • Arizona College of Osteopathic Medicine, Midwestern University, Glendale, AZ, USA MATEJ KOHUTIAR • Department of Medical Chemistry and Clinical Biochemistry, 2nd Faculty of Medicine, Charles University in Prague and Motol University Hospital, Prague, Czech Republic PIOTR KOPROWSKI • Laboratory of Intracellular Ion Channels, Nencki Institute of Experimental Biology, Warsaw, Poland JORG KOTZKA • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center at the Heinrich-Heine-University Duesseldorf, Leibniz Center for Diabetes Research, Duesseldorf, Germany; German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany JUAN C. LANDONI • Research Programs Unit, Stem Cells and Metabolism, University of Helsinki, Helsinki, Finland

xiv

Contributors

JACOB L. LÉGER • Department of Chemistry and Biochemistry, Universite de Moncton, Moncton, NB, Canada STEFAN LEHR • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center at the Heinrich-Heine-University Duesseldorf, Leibniz Center for Diabetes Research, Duesseldorf, Germany; German Center for Diabetes Research (DZD e.V.), Neuherberg, Germany GUY LENAERS • UMR CNRS 6015-INSERM U1083, MitoVasc Institute, University of Angers, Angers, France MARC LIESA • Division of Endocrinology, Department of Medicine, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA LEE ANN MACMILLAN-CROW • Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, AR, USA KIANA MAHDAVIANI • Department of Medicine, Obesity Research Center, Boston University School of Medicine, Boston, MA, USA ROLAND MALLI • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria E. M. M. MANDERS • van Leeuwenhoek Center for Advanced Microscopy, University of Amsterdam, Amsterdam, The Netherlands SHIMA MEHRVAR • University of Wisconsin-Milwaukee, Milwaukee, WI, USA NATHANAEL MILLER • Division of Endocrinology, Department of Medicine, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA; Department of Medicine, Obesity Research Center, Boston University School of Medicine, Boston, MA, USA SERGIY M. NADTOCHIY • Department of Neuroscience, University of Rochester Medical Center, Rochester, NY, USA; Department of Anesthesiology, University of Rochester Medical Center, Rochester, NY, USA MEGAN NGAI • Department of Anesthesiology, University of Rochester Medical Center, Rochester, NY, USA MARCUS F. OLIVEIRA • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil MATHEUS P. OLIVEIRA • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil ZDENA PALKOVÁ • Faculty of Science, Charles University, BIOCEV, Prague, Czech Republic DMITRI B. PAPKOVSKY • School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland GARY J. PATTI • Department of Chemistry, Washington University in St. Louis, Saint Louis, MO, USA; Department of Medicine, Washington University in St. Louis, Saint Louis, MO, USA GENNADII A. PIAVCHENKO • Orel State University, Orel, Russia; I.M. Sechenov First Moscow State Medical University (Sechenov University), Moscow, Russia NICOLAS PICHAUD • Department of Chemistry and Biochemistry, Universite de Moncton, Moncton, NB, Canada GEORGE A. PORTER JR. • Departments of Pediatrics-Division Cardiology, Pharmacology and Physiology, and Medicine (Aab Cardiovascular Research Institute), University of Rochester, Rochester, NY, USA FLAVIA RADOGNA • Laboratoire de Biologie Moléculaire et Cellulaire du Cancer, Hôpital Kirchberg, Luxembourg, Luxembourg

Contributors

MD HABIBUR RAHMAN • Department of Biomedical Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China; Centre for Novel Biomaterials, The Chinese University of Hong Kong, Hong Kong SAR, China MAHSA RANJI • Biophotonics Lab, Florida Atlantic University, Boca Raton, FL, USA PASCAL REYNIER • UMR CNRS 6015-INSERM U1083, MitoVasc Institute, University of Angers, Angers, France; Department of Biochemistry and Genetics, University Hospital of Angers, Angers, France RODIESLEY S. ROSA • Federal University of Rio de Janeiro, Institute of Medical Biochemistry Leopoldo de Meis, Rio De Janeiro, RJ, Brazil MAYUKO SEGAWA • Division of Endocrinology, Department of Medicine, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA ALEKSANDRA SE˛K • Laboratory of Intracellular Ion Channels, Nencki Institute of Experimental Biology, Warsaw, Poland; Faculty of Chemistry, University of Warsaw, Warsaw, Poland AUDREY SÉMONT • Univ. Bordeaux, INSERM, Centre de recherche Cardio- Thoracique de Bordeaux, U1045 & IHU Liryc, Electrophysiology and Heart Modeling Institute, Fondation Bordeaux Université, Bordeaux, France; CHU de Bordeaux, Bordeaux, France ORIAN S. SHIRIHAI • Division of Endocrinology, Department of Medicine, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA WILLIAM I. SIVITZ • Division of Endocrinology and Metabolism, Department of Internal Medicine, The University of Iowa, Iowa City, IA, USA R. L. SMITH • Academisch Medisch Centrum Universiteit van, Amsterdam, The Netherlands NESO SOJIC • NSysA group, Univ. Bordeaux, CNRS, INP Bordeaux, ISM, UMR 5255, Talence, France AMANDA L. SOUZA • Life Science Mass Spectrometry Division, Thermo Fisher Scientific, San Jose, CA, USA ROGER SPRINGETT • CellSpex, Kent, UK OLGA A. STELMASHCHUK • Orel State University, Orel, Russia QIAN PETER SU • Institute for Biomedical Materials & Devices (IBMD), Faculty of Science, University of Technology Sydney, Sydney, NSW, Australia; School of Biomedical Engineering, Faculty of Engineering and IT, University of Technology Sydney, Sydney, NSW, Australia ANU SUOMALAINEN • Department of Neurology, University Hospital, Helsinki, Finland; Neuroscience Center, University of Helsinki, Helsinki, Finland EMMANUEL SURANITI • NSysA group, Univ. Bordeaux, CNRS, INP Bordeaux, ISM, UMR 5255, Talence, France JULIA TOBACYK • Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, AR, USA TSUNG-LIN TSAI • Institute of Microbiology and Immunology, National Yang-Ming University, Taipei, Taiwan H. VAN DER SPEK • Molecular Biology & Microbial Food Safety, Swammerdam Institute for Life Sciences (SILS), Faculty of Science (FNWI), University of Amsterdam, Amsterdam, The Netherlands HELENA L. A. VIEIRA • CEDOC, Faculdade de Ciência Médicas/NOVA Medical School, Universidade Nova de Lisboa, Lisboa, Portugal; UCIBIO, Faculdade de Ciências e Tecnologia, Universidade Nova de Lisboa, Lisboa, Portugal; Instituto de Biologia Experimental e Tecnol ogica (iBET), Oeiras, Portugal ANDREY Y. VINOKUROV • Orel State University, Orel, Russia

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Contributors

MARKUS WALDECK-WEIERMAIR • Gottfried Schatz Research Center for Cell Signaling, Metabolism and Aging, Molecular Biology and Biochemistry, Medical University of Graz, Graz, Austria AGNIESZKA WALEWSKA • Laboratory of Intracellular Ion Channels, Nencki Institute of Experimental Biology, Warsaw, Poland LIYA WANG • Department of Anatomy, Physiology and Biochemistry, Swedish University of Agricultural Sciences, Uppsala, Sweden XIAOYONG WANG • State Key Laboratory of Pharmaceutical Biotechnology, School of Life Sciences, Nanjing University, Nanjing, P. R. China AN-CHI WEI • Graduate Institute of Biomedical Electronics and Bioinformatics, National Taiwan University, Taipei, Taiwan VOLKMAR WEISSIG • Department of Pharmaceutical Sciences and Nanocenter of Excellence, Midwestern University College of Pharmacy at Glendale, Glendale, AZ, USA DANE WOLF • Division of Endocrinology, Department of Medicine, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA; Department of Medicine, Obesity Research Center, Boston University School of Medicine, Boston, MA, USA QINRU XIAO • Department of Biomedical Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China LIPING YU • Division of Endocrinology and Metabolism, Department of Internal Medicine, The University of Iowa, Iowa City, IA, USA MAŠA ŽDRALEVIĆ • Faculty of Medicine, University of Montenegro, Podgorica, Montenegro SHIRUI ZHAO • Department of Biomedical Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China; Shun Hing Institute of Advanced Engineering, The Chinese University of Hong Kong, Hong Kong SAR, China ALEXANDER V. ZHDANOV • School of Biochemistry and Cell Biology, University College Cork, Cork, Ireland BORIS ZHIVOTOVSKY • Division of Toxicology, Institute of Environmental Medicine, Karolinska Institutet, Stockholm, Sweden; Faculty of Medicine, MV Lomonosov Moscow State University, Moscow, Russia

Chapter 1 Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations, Pitfalls, and Tomorrow Naig Gueguen, Guy Lenaers, Pascal Reynier, Volkmar Weissig, and Marvin Edeas Abstract Until recently restricted to hereditary mitochondrial diseases, mitochondrial dysfunction is now recognized as a key player and strategic factor in the pathophysiological of many human diseases, ranging from the metabolism, vascular, cardiac, and neurodegenerative diseases to cancer. Because of their participation in a myriad of cellular functions and signaling pathways, precisely identifying the cause of mitochondrial “dysfunctions” can be challenging and requires robust and controlled techniques. Initially limited to the analysis of the respiratory chain functioning, these analytical techniques now enlarge to the analyses of mitochondrial and cellular metabolism, based on metabolomic approaches. Here, we address the methods used to assay mitochondrial dysfunction, with a highlight on the techniques used in diagnosis on tissues and cells derived from patients, the information they provide, and their strength and weakness. Targeting mitochondrial dysfunction by various strategies is a huge challenge, requires robust methods of evaluation, and should be able to take into consideration the mitochondria dynamics and localization. The future of mitochondrial medicine is strongly related to a perfect comprehension of its dysfunction. Key words Mitochondrial Metabolomics

dysfunctions,

Mitochondria

evaluation,

Bioenergetics,

Devices,

Introduction The contribution of mitochondrial dysfunctions in the onset and development of many diseases has been widely studied. These dysfunctions could interfere with many cellular, metabolic, and homoeostatic functions [1]. It is therefore not surprising that, beyond primary mitochondrial disorders, mitochondrial dysfunctions now impact most areas of medical. Mitochondrial dysfunction has been found in numerous common human diseases, including metabolic diseases [2] such as obesity and diabetes, heart diseases [3], neurodegenerative diseases, such as Parkinson’s, Alzheimer’s

Volkmar Weissig and Marvin Edeas (eds.), Mitochondrial Medicine: Volume 2: Assessing Mitochondria, Methods in Molecular Biology, vol. 2276, https://doi.org/10.1007/978-1-0716-1266-8_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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[4], or Huntington’s [5], infectious diseases [6], inflammation [7], and cancer [8]. Multiple reports have shown the impact of dysfunctional mitochondria on the immune response. One of the example is the COVID-19 pandemic caused by the coronavirus (SARS-CoV-2). Many recent studies revealed that human alveolar epithelial cells with dysfunctional mitochondria displayed increased production of pro-inflammatory cytokines which were found to be increased in COVID-19 [9, 10]. Recently, we proposed that not only the intracellular mitochondria dysfunction is a consequence of COVID-19 infection formation [6, 11] but also the less explored extracellular mitochondria (specifically platelets mitochondria) may affect blood coagulation, clot, and thrombosis formation [10– 15]. These extracellular mitochondria may represent a crucial intercellular mediators and may serve as strategic therapeutic targets in COVID-19 pathogenesis [11]. Historically, mitochondrial diseases have been largely addressed from the point of view of the defect of the respiratory chain and therefore of a defect in energetic homeostasis. However, mitochondria are now recognized to perform multiple additional cellular functions beyond energy production. For example, electron transfer chain (ETC) activity regulates the NADH/NAD+ redox state through NADH oxidation by complex I, [16] which impacts to hundreds of cellular reactions [17, 18]. Mitochondria are the major source of reactive oxygen species (ROS) [19], which not only contribute to normal cell function but also are linked to increased intracellular oxidative stress. ROS production regulates different signaling pathways, from metabolic rewiring to inflammatory response or cell survival [20, 21]. Mitochondria also integrate main cellular catabolic and anabolic pathways. They host not only whole or partial components of several converging catabolic cellular processes, i.e., glucose metabolism and the tricarboxylic acid (TCA) cycle, and the β-oxidation but also several biosynthetic pathways, such as folate and sulfur metabolism or heme biosynthesis. In addition to generating reducing equivalents that feed the OXPHOS system, the TCA cycle has numerous anabolic roles, providing precursors for the lipids, proteins, carbohydrates, and nucleotides biosynthesis [22]. TCA cycle metabolites also act as signaling molecules, affecting critical cellular processes that contribute to oncogenic transformation, such as nutrient signaling, HIF1α-dependent metabolic reprogramming, or histone acetylation and demethylation by acetyl-CoA and α-ketoglutarate, respectively [23]. Mitochondria are also involved in calcium homoeostasis [24], stress response and quality control [25], and initiation of caspase-dependent apoptosis [26]. Developing precise technologies to study mitochondrial physiology is getting much more important as the prevalence of common and inherited mitochondrial disease increases. Targeting

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

mitochondrial dysfunction by various strategies is a huge challenge and need a robust methods of evaluation [27]. Here, we address the techniques commonly used in clinical diagnosis, the information that can be inferred from them, their limitations, pits, and pitfalls.

Methods to Assess Mitochondrial Dysfunctions

2.1 Basis in Bioenergetic

Principles of mitochondrial bioenergetics have been largely reviewed over years [28]; therefore, only a rapid overview will be given here. Mitochondria are surrounded by the outer (OMM) and inner (IMM) membrane. The inner membrane is largely impermeant and constitutes the major barrier between the cytosol and the mitochondrial matrix. IMM forms multiple cristae, which host the ETC complexes and the enzymes involved in ATP synthesis, the F0F1-ATP synthase, the adenine nucleotide translocator (ANT), and the inorganic phosphate (Pi) transporter (PiC). The ETC system is composed of multisubunits enzymes functionally and physically linked together, the complex I (CI, NADH ubiquinone reductase), complex II (CII, succinate ubiquinone reductase), complex III (CIII, ubiquinol cytochrome c reductase), and complex IV (CIV, cytochrome c oxidase), which transfers the electron energetic potential from NADH/NAD+ (CI) and FADH2/FAD+ (CII) to the electrochemical proton gradient known as the protonmotive force (Δp). This process involved a series of oxidoreductase reactions in which electrons flow sequentially “downhill” along the ETC from a reduced to an oxidized state, ending to molecular oxygen reduction to a water molecule, or “respiration.” The release of free energy during electron transfer drives the proton pumping across the IMM at complexes CI, CIII, and CIV, in turn producing the Δp. Other oxidoreductases of the ETC are unable of proton pumping, particularly the complex II, the glycerol phosphate dehydrogenase, and the electrontransferring flavoprotein quinone oxidoreductase (ETF) linked to fatty acid β-oxidation. Thus, ten protons are extruded for each electron pair transferred from NADH to oxygen, or six protons for each electron pair transferred from FADH2 to O2, meaning that the H+/O2 and ATP/O2 stoichiometries differ according to the initial substrates. The fueling of reducing molecules, namely NADH and FADH2, to the ETC is mainly ensured by the TCA cycle and the β-oxidation pathways. According to the anaplerotic routes which feed TCA or the relative contribution of TCA or β-oxidation, the resulting NADH/FADH2 ratio differs, ultimately modifying the efficiency of ATP synthesis. The substrate oxidation module consists of all these reactions involved in substrate metabolisms and electron transport, finally generating Δp.

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The Δp is composed of the charge (Δψm) and chemical (ΔpH) components. The energy available in Δp drives the synthesis and transport of ATP, as the protons return to the matrix through the ATP synthase [29]. The ATP synthesis module includes, in addition to the ATP synthetase, the ANT that exchange the ADP/ATP and the PiC. The Δp is central to the physiological functions of mitochondria, connecting the substrate oxidation module to the ATP synthesis. Under most circumstances, proton flux is tightly coupled to OXPHOS. However, the Δp can also be dissipated by proton leak [30], in which protons return into the matrix independently of ATP synthesis, or be used for transport processes across the IMM of metabolite, ions, or calcium [31]. The total proton entry and extrusion are exactly balanced under steady-state conditions. Forward flux through ETC complexes requires a thermodynamic disequilibrium, i.e., the free energy available from electron transfer must be greater than that required to pump protons against the Δp. The Δp variations thus “dictate” the ETC activity: any decrease in the Δp is followed by an increase in electron transfer and proton pumping. In other words, the net flux of the proton current is reflected by the rate of mitochondrial oxygen consumption. Decrease in the Δp results from either proton leak or ATP synthesis, which is regulated by the cellular ATP demand. Any alteration in one the above described processes leads to a mitochondrial dysfunction. Subsequently, we shall describe the conventional methods for dissecting these processes, and identifying the primary site of impairment, starting with the methods used in the laboratory for the biochemical diagnosis of mitochondrial diseases. A complete set of these analyses will draw a picture of the mitochondrial energy-generating system. 2.2 Analyses of Maximal ETC Activities

Evaluation of individual respiratory chain complex activities is a routine biochemical approach for the diagnosis of mitochondrial disorders. Assays to quantify CI, CII, CIII, CIV, and ATP synthase enzymatic activities are performed either on tissue, mostly muscle or liver biopsies, or on cells as primary fibroblasts or lymphocytes. They require the preparation of tissue homogenates, after the elimination of cell debris and nuclei, or cell lysates, from either fresh or frozen sample. However, frozen samples are more commonly used because fresh samples must be processed immediately after sampling. Isolated activity of each complex can be analyzed by following the oxidation/reduction of specific substrates or substrate analogs by spectrophotometry. The spectrophotometric enzyme assays are: l

NADH:ubiquinone oxidoreductase (NUR, EC 7.1.1.2, NADH oxidation followed at 340 nm) for CI.

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

Succinate:ubiquinone oxidoreductase (coupled to 2,6-Dichloroindophenol reduction followed at 600 nm) for CII (SUR, EC 1.3.5.1).

Succinate:cytochrome c oxidoreductase (CII + CIII, cytochrome c reduction at 550 nm).

Ubiquinol cytochrome c oxidoreductase for CIII (UCCR, EC 1.10.2.2, cytochrome c reduction at 550 nm).

Cytochrome c oxidase for CV (COX, EC 1.9.3.1, cytochrome c oxidation at 550 nm).

The measurement of complex V (oligomycin-sensitive ATPase) is more challenging and requires mitochondrial-enriched fractions and is often measured in cultured skin fibroblasts. Citrate synthase activity is often used for normalization of respiratory chain complex activities to mitochondrial mass. The activities normalized to CS and/or the ratios between mitochondrial enzymes give a much narrower range of normal values compared to activities expressed with respect to sample protein content, since the mitochondrial mass are highly variable among individuals and subjected to mitochondrial biogenesis regulation. While the principles of the different protocols are similar, nowadays, there is no universal assay for spectrophotometric quantification of ETC enzyme activities. According to laboratories, assays differ with respect to buffer conditions, reaction temperatures, substrate concentrations, supplementation in Ca2+ chelator (EDTA), or addition of bovine serum albumin. However, in an attempt to facilitate interlaboratory comparison of the results, some reference centers for the diagnosis of mitochondrial diseases have reevaluated and standardized their protocols [32]. Accurate enzymatic testing of tissue and cell samples can be hampered by different pitfalls. A frequent cause for problems is sample collection and handling. For optimal preservation of mitochondrial enzymes, samples must be immediately snap-frozen after collection and stored at 80 C, without any thawing until analysis. Other artifacts come from local anesthetic contamination of the sample, particularly lidocaine [32, 33], or, for studies on animals, from euthanasia procedures using CO2. Furthermore, one of the main difficulties in achieving reliable and reproducible dosages is the quality of the reagents, with strong variability between reactive references or even batches of reactive. Thus, these dosages require the inclusion of quality controls in each series, in order to validate the reaction mix. This quality is usually obtained from culture cells or animal tissue [34, 35], since the human tissues are limited. In addition, assays that determine the amounts of OXPHOS complexes, such as blue native (BN) gel electrophoresis followed by Western blot analysis, should be performed to decipher an ETC defect related to decreased catalytic activity or involving a lower enzyme quantity.

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2.3 Structural Analyses: BN-PAGE

ETC enzymes are all multimeric and encoded by both the mtDNA and nDNA, except for complex II that is fully encoded by the nuclear genome. Their assembly into a functional complex requires an intricate process assisted by chaperones [36]. Nowadays, the mechanisms of assembly for each complex are almost completely resolved [36, 37]. These assembly processes require the striking coordination of intra- and extramitochondrial transcription, translation, protein import into mitochondria, and protein folding and incorporation into assembly intermediates, up to the final complex assembly. Any disturbance in one of these processes would alter complexes assembly and compromised ETC function and energy production. Mutations that impair this assembly process are a frequent cause of mitochondrial inherited diseases [38, 39]. Complex misassembly is not only associated with an increasing number of mutations in ETC subunit genes or chaperones [40] but CI misassembly has also been recently observed in neurodegenerative diseases such as Alzheimer’s and Parkinson’s diseases [41], thus broadening the spectrum of assembly defect-associated diseases, and could be implied in the ageing process [42]. Finally, assembly of CI, III, and IV together leads to respiratory super complexes, adding a layer of complexity in the structural organization of the respiratory chain. The crystal structure of the super complex has recently been elucidated [43]. Although the pathways that lead to their formation and their function are still not completely clear [37, 44], they are now recognized as the final functional ETC unit, or “respirasome.” “Blue Native” polyacrylamide gel electrophoresis (BN-PAGE) [45] is a relatively easy approach for analyzing the assembly and abundance of OXPHOS complexes for the diagnosis of mitochondrial diseases [46, 47]. The assembly profile provides information for identifying disease-causing mutations and facilitates molecular investigation by highlighting potentially involved subunits [39, 48]. BN-PAGE is performed on isolated mitochondria or enrichedmitochondrial fractions from either tissues or cells. The protocols are described in the literature [46, 47, 49]. However, it should be stressed that results depend on the choice and concentration of the detergent and on the choice of the antibodies. After mitochondrial isolation, the mitochondrial membranes are solubilized using nonionic detergent. Because mitochondrial membranes have low cholesterol but high cardiolipin contents, [50], digitonin is preferentially used for super complexes analysis while β-dodecylmaltoside should be used for isolated complex visualization. However, both the concentration and the detergent to protein ratio must be carefully optimized to ensure that detergent concentration is above critical micelle concentration but below the detergent:protein ratio that would solubilize the complexbound cardiolipin, which are critical for their stability and activity.

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

In addition, after transfer of the electrophoretic gels for Western blotting, the choice of the antibodies will determine whether the quantification of the abundance of the fully assembled complex is privileged or the detection of assembly intermediates. In principle, any subunit can be targeted and would allow the detection of the assembled complex, provided that the epitope is accessible for the antibody. However, according to the location of the subunit within the complex and the incorporation step during the assembly process, the assembly intermediates which could be detected and their number will vary (Fig. 1). Moreover, due to the relatively low dynamic range of chemiluminescence detection, the high intensity of the signal produced by the holoenzyme could blunt other bands of lower abundance, as the assembly intermediates. To address this issue, the detection of assembly factors that specifically bind intermediates, but detach once the holoenzymes are fully assembled should be preferred. In this case, only the assembly intermediates will be visualized. An example of CI assembly analysis using antibodies targeting assembly factors is illustrated in Fig. 1. It is therefore highly recommended to master the assembly pathway of the complex of interest and the location of the different subunits and assembly factors within the complex structure/intermediates before performing a BN-PAGE analysis. However, a normal respiratory chain enzyme activity/assembly does not exclude a functional impairment of the respiratory chain functioning, which may not be detectable by enzymatic measurements or assembly analyses. 2.4 Functional Analyses: Respiration Rates

Respiration studies are an efficient way to analyze ETC and metabolic activities of cells or tissues. The last, irreversible, step of electron transfer along the ETC is the transfer catalyzed by the CIV of four electrons to a molecule of oxygen to generate two molecules of water. The coupling between electron transport (oxidation) and proton pumping within ETC and the tight coupling between oxidation and phosphorylation through the Δp (51, 52) mean that the mitochondrial respiration rate is an accurate measure of the total ETC activity and mitochondrial ATP synthesis rate. According to the experimental design, information can be gained on multiple processes required for respiration, including substrate transport into the mitochondria, reducing equivalent production by TCA cycle or beta-oxidation and electron delivery to the respiratory chain, activities of the different complexes, ATP synthesis, proton leak, and mitochondrial metabolism. Beyond metabolic analysis, O2 consumption is now also used to analyze cytotoxicity.

2.4.1 Devices

The rate of mitochondrial O2 consumption could be determined by a number of methods, the two main approaches are amperometric O2 sensors [51] and O2-dependent quenching of porphyrin-based phosphors [52]. The amperometric approach, notably developed

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Fig. 1 Detection of CI assembly intermediates by targeting the CI assembly factors. (a) Schematic representation of the modular assembly of CI The CI holoenzyme results from the sequential assembly of ten intermediates (A.I.), assisted by at least 13 CI specific assembly factors. The Q earliest intermediate is composed of the NDUFS2, NDUFS3, NDUFS7, NDUFS8, and NDUFA5, stabilized by two specific assembly factors, namely NDUFAF3 and NDUFAF4 in a 170 kDa intermediate (1). The binding of Q module to the ND1 subunit and accessory subunits forms the 237 to 283 kDa (2) intermediates named Q/PP-a, which are stacked to the inner membrane by NDUFAF5 and NDUFAF6. The P module is assembled in four distinct intermediates, two distal, named PD-a and PD-b that contain ND4 and ND5 subunits, respectively, and two proximal, the PP-a and PP-b. The central PP-b intermediate (3) is the entry point for four of the seven MT-DNA-encoded subunit, i.e., ND2, ND3, ND6, and ND4L, and is bound by six assembly factors, among which, NDUFAF1. Then, the different modules progressively combine (4, 5), forming a Q/P intermediate stacked with the assembly factor NDUFAF2 (6). In the last step of the process, the 160 kDa N module (7), constituted by the catalytic NDUFV1, NDUFV2, NDUFS1, and the accessory NDUFA2 and NDUFS4 subunits, is added. Finally, once the CI assembly is completed, all the assembly factors are released, leaving a functional holoenzyme of 980 kDa (8). Nuclear-

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

by Oroboros Instrument, has historically been the most common for in vitro and in vivo investigations of mitochondrial respiration. Nevertheless, phosphorescent probes are gaining interests since the introduction of the XF Extracellular Flux Analyzer by Seahorse Bioscience [53]. The O2K® Oxygraph

The O2K® oxygraph developed by Oroboros measures oxygen consumption by polarography with a Clark’s electrode. Briefly, oxygen diffuses through a Teflon membrane, which is permeable to uncharged gases, but not to water. A platinum/silver/KClcoupled electrode reduces oxygen and oxidizes silver, giving rise to a current, which is proportional to oxygen concentration within the physiological range of measurements. It operates in a closed chamber of 2 ml, with a stirring magnet to homogenize oxygen and biological material. The volume can be adjusted according to the cells, tissues, or mitochondria concentrations to analyze. Highresolution designs of Oroboros Instruments are optimized for high sensitivity, precision, and minimal measurement interferences (typically the detection limit is ~1 pmol/sec/ml and the quantification limit is ~2.5 pmol/s/ml, on site evaluations).

Fig. 1 (continued) encoded subunit names were shortened by omitting the leading “NDUF.” Only the main subunits are indicated. The numbers refer to the corresponding intermediates that can be detected by BN-PAGE in panel b. (b) Study of CI assembly by BN-PAGE in two control fibroblasts (Ctr), one patient cell line with a known assembly defect [48] and cells lacking MT-DNA (143B Rho0). (b1) First, CI assembly was analyzed using “classical” antibodies targeting CI structural subunits located in different intermediates, i.e., NDUFS2 (Q, left panel), NDUFB6 (PD-a, middle panel), and NDUFS1 (N, right panel). In Ctr cells, anti-NDUFS2 antibody allowed the detection of the fully assembled holoenzyme (8), while Q/P A.I. (6) accumulated in the NDUFS4 mutated cells. No CI holoenzyme or A.I. could be detected in Rho0. Further hybridization with antiNDUFB6 and NDUFS1 antibodies highlighted the presence of the Pp-b/PD-a at ~700 kDa (5) and the N module (7), respectively. (b2) A.I. were then analyzed by targeting the assembly factor NDUFAF4 (Q), NDUFAF1 (Pp-b), and NDUFAF2 (Q/P). The detection of NDUFAF1 revealed a main band at ~400 kDa which matched with the described size for the Pp-b intermediate and two higher faint bands just below and above 800 kDa that matched the Q/Pp and Q/P intermediates, respectively. As expected, these intermediates containing the MT-DNA encoded subunits were not observed in Rho0 cells. NDUFAF4 antibody detected the expected bands for Q-containing intermediates, i.e., a main one for Q at 170 kDa (1) and fainter ones for Q/Pp-a (at ~240 kDa and ~280 kDa, (2)), Q/Pp (~750 kDa, (4), and Q/P (~880 kDa, (6)). As expected, only the smallest Q intermediates, constituted only of nDNA-encoded subunits, were detected in Rho0 cells. The Pp-b intermediate was detected on the same membrane by hybridizing anti-NDUFAF1 antibody. Finally, additional hybridization with NDUFAF2 antibody clearly highlighted the accumulation of the Q/P (6) that occurred in NDUFS4 cells and highlighted different higher molecular weights up to ~1100 kDa for this intermediate. Thus, since they do not bind the CI holoenzyme, whose intense signal can blunt the presence of A.I., the targeting of the assembly factors allows a more sensitive detection of these intermediates. The two combined approaches allow the identification of seven of the assembly intermediates in addition to the holoenzyme

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This is allowed by (1) closed, air-tight reaction chambers; (2) high sensitivity of the sensors with minimal noise/signal ratio; and (3) low O2-permeable materials, which minimizes O2 backdiffusion and overestimations of respiration. In these systems, instrumental background O2 consumption (i.e., nonbiological O2 flux, which constitutes potential sources of systematic error) can be corrected for accurate determinations of O2 consumption. The device can be fully calibrated, both for 0 and 100% oxygen signals and oxygen concentration within the respiratory medium according to the actual PO2 and O2 solubility. Therefore, real-time absolute quantitative measurements of oxygen concentration and fluxes can be measured, and robust comparison of inter-assays achieved. Normalization of fluxes to protein content or cell concentration can be easily achieved by recovering the samples within the chamber at the end of the experiment. O2K® oxygraphs are a fully open system, allowing as many injections of substrates or inhibitors as needed, and reoxygenation during the experiment time-course if needed, thus enabling extended substrate–uncoupler–inhibitor–titration protocols to be applied. Each complex of the ETC can be studied independently, on the same sample, using different substrate combinations. The capability of the O2k® system has recently expanded beyond respiration to permit simultaneous fluorescence-based measurements, potentiometric measurements of H+ concentrations (i.e., pH), of ΔΨm using triphenylphosphonium or fluorescent membrane potential probes, and Ca2+, as well as amperometric measurement of nitric oxide. There are however main limitations when considering the use of Oroboros technology: First, the O2k® is not a high-throughput technology, as only two samples can be performed at one time, and it is not automated. Because analysis using substrate–uncoupler– inhibitor–titration protocols take approximately one hour to complete, large-scale analyses are time-consuming. Second, analysis requires cell dissociation and suspension. As most cell culture– based studies are conducted on adherent cells, this system shows limits when the experimental objective is to assess cellular bioenergetics in intact cultured cells. According to cell types, cell detachment from the extracellular matrix could alter metabolism [54], in any way, analyses must be performed immediately and achieved rapidly after cell detachment. Third, despite the high sensitivity, the relative high volume of the chambers (500 μl to 2 ml) imposes a nonnegligible sample quantity. Typically, 10–40 μg of isolated mitochondria, 1–2 mg permeabilized tissue or 2–6·106 cells, according to cell types, are needed for one experiment.

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . . The Seahorse® XF

The Seahorse® XF Extracellular Flux Analyzer technology is based on solid state sensor probes for detection of oxygen and H+ concentrations, residing 200 μm above the cell/sample monolayer on a multi-well microplate format. Real-time measurements of oxygen concentrations are made by isolating a small volume of about 2 μl of medium above a monolayer of samples in a “transient microchamber” within the microplate. The technology based on fluorescence quenching is quite sensitive and has been optimized to avoid signal drifting. The main advantage of Seahorse® technology relies on its multi-wells plate format and automated system, allowing highthroughput analyses. Moreover, the small chamber volume is a great benefit when limited material is available. Finally, this system represented a technical breakthrough for the field of bioenergetics by providing the first instrument with the capacity to measure in vivo O2 consumption on intact adherent cells in culture. The XF Analyzer minimizes the potential limitation of oxygen diffusion for adherent cells by transiently limiting the volume of solution in which O2 is measured to just above the monolayer of cultured cells. Moreover, the capability of the system expands beyond respiration and allows simultaneous measurements of H+ extrusion (i.e., pH), as a reflect of glycolytic rates. However, there are main limitations to consider when analyzing mitochondrial respiration using the Seahorse® Analyzer. First, as for many miniaturized setups, high-throughput methods, the device loses in accuracy what it gains in rapidity. The biosensors are calibrated for 100% oxygen, using the supplier solution, not in the respiratory medium used. The calibration procedure does not consider the real oxygen concentration within the medium (oxygen solubility within the medium, real barometric pressure during the experimental course). However, the temporary microenvironment created over the cell layer is not completely sealed from atmospheric O2; thus, as O2 concentration declines with time, O2 gradient favors diffusion from the media to the micro-chambers. O2 gradient could also favor back-diffusion from the plastic of the wells. As the device back-diffusion of oxygen is not quantified and background correction is performed relative to the “background plate noise” determined on different “control” wells (devoid of sample), this could lead to an underestimation of cellular respiration rate. Thus, this system allows quite quantitative determination of O2 variation, but it did not allow absolute quantitative measurements of O2 concentration and fluxes. Second, inherent to the technologies based on fluorescence quenching is also the quite high variability between the different acquisitions (intra-assay variability (intraplate, inter-sensors) and more strikingly, inter-plates variability) [55]. This limits the comparison between different experiments. Third, the Seahorse® setup is not an open system, and only four ports surrounding the sensor are available that must be loaded

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before starting the analysis. This strongly limits the ETC substrates/inhibitors titration protocols. It is essential to perform robust optimization steps prior to any respiratory assay: the optimal concentration of each injectable reagent must be checked for each experimental condition. Care must be also taken not to overload the wells, which can cause O2 to quickly become depleted under phosphorylating conditions, leading to hypoxia. Finally, this device requires that samples adhere to the bottom of the well, which corresponds to an advantage when working on adherent cells, but could require coating of the samples in other conditions. To resume, when accurate measurement of O2 consumption and high versatility is aimed, as for diagnosis purpose, highresolution O2K® respirometry system should be used. However, if high-throughput dosages are needed as for the screening of drugs, or when the sample quantities are limited, the Seahorse® offers a much higher throughput platform than the traditional Clark electrode-based systems. 2.4.2 Permeabilized Tissues and Cells

Within tissues or cells, mitochondria are not accessible for many substrates and inhibitors, and the large catabolic processes that must be considered when dissecting ETC activity and mitochondrial metabolism greatly complicated the understanding of the results. Therefore, Isolation of mitochondria through tissue/cell culture homogenization and differential centrifugations is routinely used for assessment of mitochondrial respiration [56, 57]. Isolated mitochondria remain one of best approaches for studying mitochondrial bioenergetics free from the influence of cellular factors like the cytoskeleton, endoplasmic reticulum, cellular ATPases, together with a strict control on substrate supplies. It also allows the studies of distinct subcellular mitochondria, such as subsarcolemmal and intermyofibrillar mitochondria of skeletal muscle, which display different functional properties [58]. However, the disadvantages of isolated mitochondria include: 1. The disruptions of mitochondrial structure [59], of mitochondrial network, mitochondria–endoplasmic reticulum and mitochondria–cytoskeleton interactions, which may impact function [60–62]. 2. The purification process also bias the mitochondrial composition, by selecting high-density mitochondria, while discarding less dense ones during differential centrifugation steps [63]. 3. The purification process requires relatively large sample sizes (roughly 20 106 cells, or 100–150 mg wet weight of tissue) to obtain relevant yields. 4. The loss of micro-compartmentalization and metabolic channeling [64, 65].

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

An alternative approach when the control of substrate supply is needed is the use of permeabilized tissues, mostly permeabilized muscle fibers, or cells using low concentration of plasma-selective detergents [66]. Saponin and digitonin, a saponin derivate, are mostly selective for cholesterol lipids, enriched in the plasma membrane. Therefore, the use of saponin (50–100 μg/ml) for tissue permeabilization or digitonin (10–30 μg/million cells) for cell permeabilization leaves intact all intracellular structures, including mitochondria. Permeabilized cell and skinned fiber techniques have several advantages for studies of mitochondrial function: 1. very small tissue samples are required; 2. all mitochondria can be investigated; 3. more importantly, mitochondria are studied in their natural surroundings, as the mitochondrial network is maintained as well as the potential interactions between mitochondria and other subcellular structures and organelles. However, the mitochondrial integrity after sample preparations must be carefully controlled by checking the absence of any stimulatory effect of cytochrome c, witnessing the maintenance of the mitochondrial membranes’ integrity. The classical respiration rate experiments to determine mitochondrial bioenergetic function were defined by Chance and Williams [67]. Substrate is added in the respiratory medium (defined as “state 2,” no ADP), followed by addition of ADP, allowing the ATP synthase to function. This induces a drop in Δp and thus accelerates the electron transport and proton pumping (“state 3”). On permeabilized fibers or cells, the presence of cellular ATPase activity maintains a high ADP/ATP ratio through constant ATP hydrolysis and subsequent ATP recycling by mitochondrial synthesis. State 4o is achieved by adding oligomycin, the ATP synthase inhibitor, to inhibit ATP synthesis, increasing back the Δp resulting in slowing down the respiration rate. Although state 2 and state 4 respiration are experimentally quite equivalent, state 4 is more conveniently used when referring to respiration under basal, non-phosphorylating conditions. Then, addition of a protonophore, such as FCCP (carbonyl cyanide-p-trifluoromethoxyphenylhydrazone), gives the uncoupled respiration (“state 3u”). State 3u is controlled exclusively by substrate oxidation module (including substrate uptake, activity of the NADH/FADH2 producing pathway, e.g., TCA cycle, activity of the ETC complexes, pool sizes of ubiquinone, and cytochrome c). Inhibition of any of these processes will decrease the state 3u rate, but due to the presence of controlled, unlimited substrates, state 3u mainly reflect the maximal ETC capacity for a given substrate, even if the TCA cycle activity could be sometimes rate-limiting.

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State 3 (ADP) is controlled, depending on the tissue and conditions, by both the ATP synthesis module (ANT, PiC, and ATP synthase) and substrate oxidation. Indeed, in some mitochondria, such as those from brown fat or liver, the ATP synthase activity is limited and displays a striking control over the ETC activity, meaning that the maximal ETC activity can only be reached in uncoupling conditions [68]. Comparing state 3u and state 3, when ADP is added in saturating concentration, will thus provide insight into the control exerted by the ATP synthesis module on the substrate oxidation one. State 4 is controlled predominantly by the proton leak and to a small extent by the activity of substrate transport. In optimal condition, mitochondria maintain a high Δp which restricts proton pumping and thus electron transport leading to slowdown of the respiration rate. An increased state 4o rate for a given substrate would indicate altered proton leak. Importantly, the specific subset of complexes and dehydrogenases engaged by these assays depends on the substrates provided. These respiratory states can be sequentially measured in the presence of different combinations of mitochondrial substrates, allowing multiple metabolic pathways or respiratory complexes to be probed. A typical titration protocol used in diagnosis pipelines, using substrates of CI, CI + CII, CII, and IV, is as followed (Fig. 2): First, state 2 (non-phosphorylating) respiration is initiated after adding pyruvate and malate. Second, the state 3 respirations are detailed for the different complexes: the CI-linked maximal phosphorylating respiration is stimulated by saturating ADP concentration (1.5 mM). Succinate (10 mM) is added to measure the combined CI and CII-linked respiration with convergent CI + II electron flow into the Q-junction corresponding to the maximal stimulated phosphorylating respiration (OXPHOS capacity). Rotenone is then used to inhibit CI activity and thus to obtain the maximal CII-linked phosphorylating respiration. Third, oligomycin is used to inhibit F0F1-ATP synthase and state 4o measurement. FCCP is sequentially added to uncouple mitochondria and measure the CII-linked respiration in state 3u. Antimycin A (2 μg/ml) is used to inhibit complex III and check for the non-mitochondrial oxidation. Finally, ascorbate + TMPD, the artificial complex IV substrates, allow the measurement of maximal COX-driven respiration rate, after inhibition of complex IV by potassium cyanide and subtraction of this nonspecific oxidation. The respiratory control ratio (RCR), defined as the quotient of maximal state 3 to state 4 respirations, is often used as an index of mitochondrial coupling of oxidation to phosphorylation for a given substrate combination. The RCR indeed integrates the increase in

Mitochondrial Dysfunction in Mitochondrial Medicine: Current Limitations. . .

Fig. 2 Typical analyze traces of O2 consumption on permeabilized fibers. (a) Segmental analysis of ETC function using sequential substrates injections. Mitochondrial oxygen consumption measurements were performed at 37 C and atmospheric pressure using a high-resolution oxygraph (O2K, Oroboros Instrument, Innsbruck, Austria). Respiration rates on permeabilized fibers using 50 μg/ml saponin (30 minutes, 4 C) were measured in respiratory buffer (10 mM KH2PO4, 300 mM mannitol, 10 mM KCl, 5 mM MgCl2, 0.5 mM EGTA, and 1 mg/ml serum albumin bovine, pH 7.2) using substrates of CI, CI + CII, and CII as followed: first, state 2 (non-phosphorylating) respiration was measured after adding 2.5 mM pyruvate and 5 mM malate. Then, the CI-linked maximal phosphorylating respiration was stimulated by saturating ADP concentration (1.5 mM) and 3 mM NAD+, added to avoid TCA limitation by NAD+ availability. CI-linked maximal phosphorylating respiration was further stimulated with 5 mM glutamate, to check for any limitation of substrate availability by PDH activity. Succinate (10 mM) was then added to measure the combined CI and CII-linked respiration with convergent CI + II electron flow into the Q-junction corresponding to the maximal stimulated phosphorylating respiration (OXPHOS capacity). Rotenone (5 μM) was used to inhibit CI activity and thus to obtain the maximal CII-linked respiration. Thirdly, oligomycin (F0F1-ATP synthase inhibitor, 4 μg/ml) and FCCP (carbonyl cyanide-p-trifluoromethoxyphenylhydrazone, a mitochondrial uncoupler, 1 μM) were sequentially added to ensure that the cells were fully permeabilized. Antimycin A addition (2 μg/ml) was used to check for the non-mitochondrial oxidation. CIV maximal respiration was induced with 4 mM ascorbate and 0.3 mM TMPD,