Mitochondria: Methods and Protocols (Methods in Molecular Biology, 2497) 1071623087, 9781071623084

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Methods in Molecular Biology 2497

Namrata Tomar Editor

Mitochondria Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Mitochondria Methods and Protocols

Edited by

Namrata Tomar Department of Biomedical Engineering, Medical College of Wisconsin, Milwaukee, WI, USA

Editor Namrata Tomar Department of Biomedical Engineering Medical College of Wisconsin Milwaukee, WI, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2308-4 ISBN 978-1-0716-2309-1 (eBook) https://doi.org/10.1007/978-1-0716-2309-1 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, Corrected Publication 2022 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. The editors would like to thank Jasiel Strubbe and Yan Levitsky from the Department of Physiology at Michigan State University for the cover art based on cryoEM images of isolated mitochondria. 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 are fascinating organelles that regulate critical cellular processes and play central roles in cell metabolism, energy production, the regulation of reactive oxygen species (ROS) emission and signaling, calcium homeostasis, and the regulation of apoptosis and energy-sensitive signaling pathways. Current practices in mitochondrial biology research include exploring basic mitochondrial biology and dysfunctions in human diseases. Investigating mitochondrial function plays a great role to understand aging, neurodegeneration, diabetes, heart failure, and inherited mitochondrial diseases. For that purpose, the book represents a set of comprehensive protocols and methodologies to assess the mitochondrial bioenergetics and dynamics in different tissues and cells involving health and pathological states. It represents updated techniques and protocols to understand complex integrated relationships among mitochondrial oxygen consumption (OCR), calcium (Ca2+) uptake and handling, ROS emission, and membrane potential. More recently, computational and mathematical models have come up that explain the mechanism and regulation of mitochondrial energy metabolism in health and pathology. This book also includes a detailed computational model to understand mitochondrial oxidative stress. The aim of the present book is therefore to bring together basic and advanced research, key experiments and protocols, and new findings related to all aspects of mitochondrial biology in healthy and diseased states. This book consists of 27 chapters that cover diverse protocols to assess mitochondrial biology.

Content and General Outline of the Book Chapter 1 describes a novel approach to sequentially assess electron flow through all respiratory complexes in permeabilized and intact cells by respirometry. This chapter also describes a highly sensitive and fast method to assess ΔΨm and NADH generation in live cells using plate reader assays. The combined method is an inexpensive and fast method for the determination of three major readouts of mitochondrial function. Chapter 2 discusses a protocol to calculate both mitochondrial membrane potential (ΔΨM) and plasma membrane potential (ΔΨP) in absolute millivolts in intact single cells, or in populations of adherent, cultured cells. This assay is optimized for 96-well microplate setup and is compatible with wide field, confocal, or two-photon microscopy. Chapter 3 describes an assay to evaluate the protective effects of rotenone or other potential inhibitors of the complex I of mitochondrial respiratory chain against acute ischemia-induced injuries in the brain. Chapter 4 discusses the methods followed in performing histoenzymology of mitochondrial complex II and III activity in the brain. The chapter also sheds light on the precautions in order to yield significant results. Chapter 5 describes an integrated toolbox, i.e., Mitotoolbox to simultaneous analyze the oxygen consumption rates (OCRs) and superoxide production on a single mitochondrial preparation. Chapter 6 provides a detailed protocol for measuring mitochondrial Ca2+ handling in the isolated functionally intact mitochondria from cardiac tissue of the guinea pig.

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Chapter 7 presents a protocol using Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) to analyze the structures of mitochondrial respiratory chain which include the preparation of the tissue samples, isolation of mitochondrial membrane proteins, and analysis of respiratory complexes. Chapter 8 describes an optimized approach to carry out multi-well measurement of tissue-specific O2 consumption using the Agilent Seahorse XFe96 analyzer on mouse brain and muscles. Protocols include the preparation of the tissue samples, isolation of mitochondria, and analysis of their functions; and preparation and optimization of the reagents and samples. Chapter 9 provides a detailed protocol to describe the mitochondrial Ca2+ retention capacity (CRC)/swelling assay and discusses how varying concentrations of mitochondria and Ca2+ added to the system affect the assay in a dose-dependent manner. Chapter 10 explores two measurements to delineate the mitochondrial function, i.e., ATP production and O2 consumption. Aerobic ATP production is quantified by phosphorus magnetic resonance spectroscopy (31PMRS) in vivo. Mitochondrial O2 consumption is quantified by O2 polarography. Chapter 11 investigates the interactions between isolated mitochondria and arsenicals. Given the ubiquitous presence of mitochondria and its role in cellular metabolism, arsenicalmitochondrial interactions may be of clinical importance that may reveal the mechanism of disease curation. Chapter 12 provides a comprehensive protocol to measure OCR and their coupling to ATP production in intact and permeabilized cells, as well as in mitochondria isolated from tissues. Chapter 13 provides two protocols to monitor Doxorubicin-induced alterations in mitochondrial morphology and respiration in isolated primary neonatal rat cardiomyocytes. This kind of study is essential as cardiac toxicity limits effective and subsequent use of DOX in chemotherapy regimens in pediatric, adult, and recurrent cancer patients. Chapter 14 describes a method to create yeast models of mitochondrial MT-ATP6 gene mutations detected in patients, to determine how the mutation impacts oxidative phosphorylation (OXPHOS). Chapter 15 describes a protocol to characterize the mitochondrial defects associated with mutations in mitochondrial tRNA genes using yeast Saccharomyces cerevisiae mutants, bearing human equivalent pathogenic mutations. This method can distinguish different protein synthesis profiles in the mitochondrial tRNA mutants. Chapter 16 provides details to engineer a yeast strain expressing a new type of GFP called Bi-Genomic Mitochondrial-Split-GFP (BiG Mito-Split-GFP) to overcome the technical difficulty caused by GFP tagging that prevents the detection of the mitochondrial echoform due to the fluorescence emission of the cytosolic isoform. Its strength lies in the mitochondrial localization of a given protein using a microscopy observation. Chapter 17 explains a fluorescence lifetime imaging microscopy (FLIM) strategy to image mitochondrial metabolic profiles in lymphocytes as they go through changes in metabolic activity. This novel imaging strategy offers a reliable tool to study changes in mitochondrial metabolism. Chapter 18 describes experimental procedures used to evaluate different parameters including mitochondrial morphology, adenosine triphosphate (ATP) levels, ROS levels, and mitophagy. Together these parameters can help to investigate mitochondria functions upon infection.

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Chapter 19 describes the simultaneous measurement of respiratory rates of isolated mouse heart mitochondria and the redox state of their Q pool using a custom-made combination of a Clark-type oxygen electrode and a Q electrode. Chapter 20 utilizes alternative oxidase (AOX), which bypasses the cytochrome pathway of the ETC when blocked to test the assumption that ETC is both an O2 sensor and a main consumer of O2 and the ETC itself adapts its electron flux to O2 availability. Chapter 21 describes the measurement of NADH and FAD+ levels in intact cells using fluorescence microscopy. The calibration of the NADH and FAD2+ fluorescence signals is a crucial factor in accurately assessing mitochondrial function and redox status. Chapter 22 describes a protocol to perform live imaging of cells incubated with TMRM and record dynamic changes using confocal laser scanning microscopy with the application of live time-series program. Chapter 23 explains a methodology that includes fluorescence and bioluminescence measurements of mitochondrial Ca2+ dynamically in both isolated mitochondria and intact cells. Mitochondrial Ca2+ uptake or mitochondrial Ca2+ efflux can be pharmacologically targeted to examine the effects on mitochondrial Ca2+ dynamics using these techniques. Chapter 24 describes a quick and reliable assay using dihydroethidium (DHE) and MitoSOX to measure cytosolic superoxide and mitochondrial superoxide production, respectively, in intact adherent cells by fluorescence microplate readers. Chapter 25 describes the application of Blue Native polyacrylamide gel electrophoresis (BN-PAGE) to analyze the organization and activity of oxidative phosphorylation (OXPHOS) complexes from cultured skin fibroblasts. Chapter 26 presents a comprehensive protocol to multiplex the Seahorse XFe24 analyzer with ImageXpress® Nano high content imaging microscope to provide a comprehensive yet rigorous profile of bioenergetics and its correlation to neuronal function. Both of these techniques work on multi-well formats. Chapter 27 details the equations behind a computational model of the mitochondria and presents simulations that validate the model against experimental data as well as model predictions from simulations. Milwaukee, WI, USA

Namrata Tomar

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

1 Fast Determination of Mitochondrial Metabolism and Respiratory Complex Activity in Permeabilized and Intact Cells . . . . . . . . . . . . . . . . . . . . . . . . . Kareem A. Heslop, Amandine Rovini, Monika Gooz, and Eduardo N. Maldonado 2 Unbiased Millivolts Assay of Mitochondrial Membrane Potential in Intact Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chad A. Lerner and Akos A. Gerencser 3 Rotenone Decreases Ischemia-Induced Injury by Inhibiting Mitochondrial Permeability Transition: A Study in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ramune Morkuniene, Evelina Rekuviene, and Dalia M. Kopustinskiene 4 Assessment of Mitochondrial Complex II and III Activity in Brain Sections: A Histoenzymological Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rubina Roy, Rajib Paul, Pallab Bhattacharya, and Anupom Borah 5 Measurement of Mitochondrial (Dys)Function in Cellular Systems Using Electron Paramagnetic Resonance (EPR): Oxygen Consumption Rate and Superoxide Production. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Donatienne d’Hose and Bernard Gallez 6 Mitochondrial Calcium Handling in Isolated Mitochondria from a Guinea Pig Heart . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jyotsna Mishra and Amadou K. S. Camara 7 Characterizing the Electron Transport Chain: Structural Approach . . . . . . . . . . . Ting Liang, Janice Deng, Bijaya Nayak, Xin Zou, Yuji Ikeno, and Yidong Bai 8 Characterizing the Electron Transport Chain: Functional Approach Using Extracellular Flux Analyzer on Mouse Tissue Samples . . . . . . . . . . . . . . . . . Ting Liang, Jay Dunn, Xin Zou, Bijaya Nayak, Yuji Ikeno, Lihong Fan, and Yidong Bai 9 Simultaneous Acquisition of Mitochondrial Calcium Retention Capacity and Swelling to Measure Permeability Transition Sensitivity . . . . . . . . . . . . . . . . . . Arielys M. Mendoza and Jason Karch 10 Measuring Mitochondrial Function: From Organelle to Organism . . . . . . . . . . . . Matthew T. Lewis, Yan Levitsky, Jason N. Bazil, and Robert W. Wiseman 11 Mitochondrial Toxicity of Organic Arsenicals. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yu-Jiao Liu and Yi Liu

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Measurements of Mitochondrial Respiration in Intact Cells, Permeabilized Cells, and Isolated Tissue Mitochondria Using the Seahorse XF Analyzer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jessica Pfleger Monitoring Mitochondrial Morphology and Respiration in Doxorubicin-Induced Cardiomyopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chowdhury S. Abdullah, Richa Aishwarya, Mahboob Morshed, Naznin Sultana Remex, Sumitra Miriyala, Manikandan Panchatcharam, and Md. Shenuarin Bhuiyan Creation of Yeast Models for Evaluating the Pathogenicity of Mutations in the Human Mitochondrial Gene MT-ATP6 and Discovering Therapeutic Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . De´borah Tribouillard-Tanvier, Alain Dautant, Franc¸ois Godard, Camille Charles, Chiranjit Panja, Jean-Paul di Rago, and Roza Kucharczyk In Vivo Analysis of Mitochondrial Protein Synthesis in Saccharomyces cerevisiae Mitochondrial tRNA Mutants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arianna Montanari Visualizing Mitochondrial Importability of a Protein Using the Yeast Bi-Genomic Mitochondrial-Split-GFP Strain and an Ordinary Fluorescence Microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marine Hemmerle, Bruno Senger, Jean-Paul di Rago, Roza Kucharczyk, and Hubert D. Becker Analysis of Mitochondrial Performance in Lymphocytes Using Fluorescent Lifetime Imaging Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meha Patel, Javier Manzella-Lapeira, and Munir Akkaya Monitoring Mitochondrial Perturbations During Infection . . . . . . . . . . . . . . . . . . Varnesh Tiku and Man-Wah Tan Measuring the Mitochondrial Ubiquinone (Q) Pool Redox State in Isolated Respiring Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marten Szibor, Estelle Heyne, Carlo Viscomi, and Anthony L. Moore Experimental Setup for Investigation of Acute Mitochondrial Oxygen Sensing in Primary Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fenja Knoepp, Norbert Weissmann, Natascha Sommer, and Marten Szibor Assessing the Redox Status of Mitochondria Through the NADH/FAD2+ Ratio in Intact Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Haoyu Chi, Gauri Bhosale, and Michael R. Duchen Monitoring Mitochondrial Membrane Potential in Live Cells Using Time-Lapse Fluorescence Imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gabriel Esteban Valdebenito and Michael R. Duchen Investigating Mitochondrial Ca2+ Dynamics in Isolated Mitochondria and Intact Cells: Application of Fluorescent Dyes and Genetic Reporters . . . . . . Gauri Bhosale and Michael R. Duchen

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A Plate Reader-Based Measurement of the Cellular ROS Production Using Dihydroethidium and MitoSOX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chih-Yao Chung and Michael R. Duchen 25 Analysis of Organization and Activity of Mitochondrial Respiratory Chain Complexes in Primary Fibroblasts Using Blue Native PAGE . . . . . . . . . . . Kritarth Singh and Michael R. Duchen 26 Multiplexing Seahorse XFe24 and ImageXpress® Nano Platforms for Comprehensive Evaluation of Mitochondrial Bioenergetic Profile and Neuronal Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Smijin K. Soman, Maryann Swain, and Ruben K. Dagda 27 Computational Modeling of Mitochondria to Understand the Dynamics of Oxidative Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rashmi Kumar and Mohsin S. Jafri Correction to: Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

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Contributors CHOWDHURY S. ABDULLAH • Department of Pathology and Translational Pathobiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA RICHA AISHWARYA • Department of Molecular and Cellular Physiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA MUNIR AKKAYA • Laboratory of Immunogenetics, National Institutes of Allergy and Infectious Diseases, NIH, Rockville, MD, USA; Division of Rheumatology and Immunology, Department of Internal Medicine, The Ohio State University, College of Medicine, Columbus, OH, USA; Department of Microbial Infection and Immunity, The Ohio State University College of Medicine, Columbus, OH, USA; Pelotonia Institute for Immuno-Oncology, The Ohio State University College of Medicine, Columbus, OH, USA DAUTANT ALAIN • Univ. Bordeaux, CNRS, IBGC, UMR 5095, Bordeaux, France YIDONG BAI • Department of Cell Systems and Anatomy, University of Texas Health San Antonio, San Antonio, TX, USA JASON N. BAZIL • Department of Physiology, Michigan State University, East Lansing, MI, USA HUBERT D. BECKER • Universite´ de Strasbourg, CNRS, Ge´ne´tique Mole´culaire, Ge´nomique, Microbiologie, UMR 7156, Strasbourg Cedex, France PALLAB BHATTACHARYA • Department of Pharmacology and Toxicology, National Institute of Pharmaceutical Education and Research (NIPER), Gandhinagar, Gujarat, India GAURI BHOSALE • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK MD. SHENUARIN BHUIYAN • Department of Pathology and Translational Pathobiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA; Department of Molecular and Cellular Physiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA ANUPOM BORAH • Cellular and Molecular Neurobiology Laboratory, Department of Life Science and Bioinformatics, Assam University, Silchar, Assam, India AMADOU K. S. CAMARA • Department of Anesthesiology and Anesthesia Research, Medical College of Wisconsin, Milwaukee, WI, USA CAMILLE CHARLES • Univ. Bordeaux, CNRS, IBGC, UMR 5095, Bordeaux, France HAOYU CHI • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK PANJA CHIRANJIT • Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland CHIH-YAO CHUNG • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK DONATIENNE D’HOSE • Biomedical Magnetic Resonance, Louvain Drug Research Institute (LDRI), Universite´ catholique de Louvain (UCLouvain), Brussels, Belgium RUBEN K. DAGDA • Department of Pharmacology, University of Nevada, Reno School of Medicine, Reno, NV, USA TRIBOUILLARD-TANVIER DE´BORAH • Univ. Bordeaux, CNRS, IBGC, UMR 5095, Bordeaux, France; INSERM, Paris, France

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JANICE DENG • Department of Cell Systems and Anatomy, University of Texas Health San Antonio, San Antonio, TX, USA JEAN-PAUL DI RAGO • Institut de Biochimie et Ge´ne´tique Cellulaires, CNRS UMR5095, Universite´ de Bordeaux, Bordeaux cedex, France MICHAEL R. DUCHEN • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK JAY DUNN • Agilent Technologies, Inc, Santa Clara, CA, USA LIHONG FAN • Department of Respiratory Medicine, Shanghai 10th People’s Hospital, Tongji University, Shanghai, China GODARD FRANC¸OIS • Univ. Bordeaux, CNRS, IBGC, UMR 5095, Bordeaux, France BERNARD GALLEZ • Biomedical Magnetic Resonance, Louvain Drug Research Institute (LDRI), Universite´ catholique de Louvain (UCLouvain), Brussels, Belgium AKOS A. GERENCSER • Buck Institute for Research on Aging, Novato, CA, USA; Image Analyst Software, Novato, CA, USA MONIKA GOOZ • Department of Drug Discovery & Biomedical Sciences, Medical University of South Carolina, Charleston, SC, USA MARINE HEMMERLE • Universite´ de Strasbourg, CNRS, Ge´ne´tique Mole´culaire, Ge´nomique, Microbiologie, UMR 7156, Strasbourg Cedex, France KAREEM A. HESLOP • Department of Drug Discovery & Biomedical Sciences, Medical University of South Carolina, Charleston, SC, USA ESTELLE HEYNE • Department of Cardiothoracic Surgery, Jena University Hospital, Center for Sepsis Control and Care (CSCC), Jena, Germany YUJI IKENO • Department of Pathology, Barshop Institute of Longevity and Aging Research, University of Texas Health San Antonio, and Geriatric Research Education and Clinical Center (GRECC), Audie L. Murphy VA Hospital, South Texas Veterans Health Care System, San Antonio, TX, USA MOHSIN S. JAFRI • School of Systems Biology, George Mason University, Fairfax, VA, USA; Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, MD, USA SMIJIN K. SOMAN • Department of Pharmacology, University of Nevada, Reno School of Medicine, Reno, NV, USA JASON KARCH • Department of Molecular Physiology and Biophysics, Baylor College of Medicine, Houston, TX, USA; Cardiovascular Research Institute, Baylor College of Medicine, Houston, TX, USA FENJA KNOEPP • Excellence Cluster Cardio-Pulmonary Institute (CPI), University of Giessen and Marburg Lung Center (UGMLC), Member of the German Center for Lung Research (DZL), Justus-Liebig-University, Giessen, Germany DALIA M. KOPUSTINSKIENE • Institute of Pharmaceutical Technologies, Medical Academy, Lithuanian University of Health Sciences, Kaunas, Lithuania ROZA KUCHARCZYK • Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland RASHMI KUMAR • School of Systems Biology, George Mason University, Fairfax, VA, USA CHAD A. LERNER • Buck Institute for Research on Aging, Novato, CA, USA; Image Analyst Software, Novato, CA, USA YAN LEVITSKY • Department of Physiology, Michigan State University, East Lansing, MI, USA

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MATTHEW T. LEWIS • Department of Internal Medicine, University of Utah, Salt Lake City, UT, USA; Geriatric Research, Education, and Clinical Center, VA Medical Center, Salt Lake City, UT, USA TING LIANG • Key Laboratory of Laboratory Medicine, Ministry of Education, Zhejiang Provincial Key Laboratory of Medical Genetics, College of Laboratory Medicine and Life Sciences, Wenzhou Medical University, Wenzhou, Zhejiang, China; Department of Cell Systems and Anatomy, University of Texas Health San Antonio, San Antonio, TX, USA YI LIU • State Key Laboratory of Membrane Separation and Membrane Process, School of Chemistry, Tiangong University, Tianjin, P. R. China YU-JIAO LIU • State Key Laboratory of Membrane Separation and Membrane Process, School of Chemistry, Tiangong University, Tianjin, P. R. China EDUARDO N. MALDONADO • Department of Drug Discovery & Biomedical Sciences, Medical University of South Carolina, Charleston, SC, USA; Hollings Cancer Center, Medical University of South Carolina, Charleston, SC, USA JAVIER MANZELLA-LAPEIRA • Laboratory of Immunogenetics, National Institutes of Allergy and Infectious Diseases, NIH, Rockville, MD, USA ARIELYS M. MENDOZA • Department of Molecular Physiology and Biophysics, Baylor College of Medicine, Houston, TX, USA SUMITRA MIRIYALA • Department of Cellular Biology and Anatomy, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA JYOTSNA MISHRA • Department of Anesthesiology and Anesthesia Research, Medical College of Wisconsin, Milwaukee, WI, USA ARIANNA MONTANARI • Department of Biology and Biotechnologies “Charles Darwin”, Sapienza University of Rome, Rome, Italy ANTHONY L. MOORE • Biochemistry & Biomedicine, School of Life Sciences, University of Sussex, Brighton, UK RAMUNE MORKUNIENE • Neuroscience Institute, Medical Academy, Lithuanian University of Health Sciences, Kaunas, Lithuania; Department of Drug Chemistry, Faculty of Pharmacy, Medical Academy, Lithuanian University of Health Sciences, Kaunas, Lithuania MAHBOOB MORSHED • Department of Pathology and Translational Pathobiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA BIJAYA NAYAK • Department of Cell Systems and Anatomy, University of Texas Health San Antonio, San Antonio, TX, USA MANIKANDAN PANCHATCHARAM • Department of Cellular Biology and Anatomy, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA MEHA PATEL • Laboratory of Immunogenetics, National Institutes of Allergy and Infectious Diseases, NIH, Rockville, MD, USA RAJIB PAUL • Department of Zoology, Pandit Deendayal Upadhyaya Adarsha Mahavidyalaya (PDUAM), Karimganj, Assam, India JESSICA PFLEGER • Center for Vascular and Heart Research, Fralin Biomedical Research Institute at Virginia Tech Carilion, Roanoke, VA, USA; Department of Biological Sciences, Virginia Polytechnic Institute and State University, Blacksburg, VA, USA EVELINA REKUVIENE • Department of Biochemistry, Faculty of Medicine, Medical Academy, Lithuanian University of Health Sciences, Kaunas, Lithuania

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NAZNIN SULTANA REMEX • Department of Molecular and Cellular Physiology, Louisiana State University Health Sciences Center-Shreveport, Shreveport, LA, USA AMANDINE ROVINI • Department of Drug Discovery & Biomedical Sciences, Medical University of South Carolina, Charleston, SC, USA RUBINA ROY • Cellular and Molecular Neurobiology Laboratory, Department of Life Science and Bioinformatics, Assam University, Silchar, Assam, India BRUNO SENGER • Universite´ de Strasbourg, CNRS, Ge´ne´tique Mole´culaire, Ge´nomique, Microbiologie, UMR 7156, Strasbourg Cedex, France KRITARTH SINGH • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK NATASCHA SOMMER • Excellence Cluster Cardio-Pulmonary Institute (CPI), University of Giessen and Marburg Lung Center (UGMLC), Member of the German Center for Lung Research (DZL), Justus-Liebig-University, Giessen, Germany MARYANN SWAIN • Department of Pharmacology, University of Nevada, Reno School of Medicine, Reno, NV, USA MARTEN SZIBOR • Faculty of Medicine and Health Technology, Tampere University, Tampere, Finland; Department of Cardiothoracic Surgery, Jena University Hospital, Center for Sepsis Control and Care (CSCC), Jena, Germany MAN-WAH TAN • Department of Infectious Diseases, Genentech Inc., South San Francisco, CA, USA VARNESH TIKU • Department of Infectious Diseases, Genentech Inc., South San Francisco, CA, USA GABRIEL ESTEBAN VALDEBENITO • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, UCL, London, UK CARLO VISCOMI • Department of Biomedical Sciences, University of Padova, Padova, Italy NORBERT WEISSMANN • Excellence Cluster Cardio-Pulmonary Institute (CPI), University of Giessen and Marburg Lung Center (UGMLC), Member of the German Center for Lung Research (DZL), Justus-Liebig-University, Giessen, Germany ROBERT W. WISEMAN • Department of Physiology, Michigan State University, East Lansing, MI, USA; Department of Radiology, Michigan State University, East Lansing, MI, USA XIN ZOU • Department of Pulmonary and Critical Care Medicine, Longyuan First Affiliated Hospital of Fujian Medical University, Fuzhou, Fujian, China

Chapter 1 Fast Determination of Mitochondrial Metabolism and Respiratory Complex Activity in Permeabilized and Intact Cells Kareem A. Heslop, Amandine Rovini, Monika Gooz, and Eduardo N. Maldonado Abstract Assessment of mitochondrial metabolism is multidimensional and time consuming, usually requiring specific training. Respiration, NADH generation, and mitochondrial membrane potential (ΔΨm) are dynamic readouts of the metabolism and bioenergetics of mitochondria. Methodologies available to determine functional parameters in isolated mitochondria and permeabilized cells are sometimes of limited use or inapplicable to studies in live cells. In particular, the sequential assessment of the activity of each complex in the electron transport chain has not been reported in intact cells. Here, we describe a novel approach to sequentially assess electron flow through all respiratory complexes in permeabilized and intact cells by respirometry. We also describe a highly sensitive and fast method to assess ΔΨm and NADH generation in live cells using plate reader assays. Thus, our combined method allows a relatively inexpensive and fast determination of three major readouts of mitochondrial function in a few hours, using equipment that is frequently available in many laboratories worldwide. Key words Electron transport chain, Mitochondria, Mitochondrial membrane potential, Mitochondrial metabolism, NADH, Oxygen consumption, Respiratory complex, TMRM, Warburg Metabolism

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Introduction Mitochondria are the major cellular source of ATP in non-proliferating cells, and relatively minor contributors to ATP production in several cancers that display a Warburg phenotype [1– 3]. In addition, mitochondria are essential for redox and ion homeostasis, biosynthesis of macromolecules, generation of reactive oxygen species, cellular signaling, and apoptosis [4–6]. Respiratory substrates are catabolized in the Krebs cycle to generate NADH and FADH2 that are donors of electrons entering the electron transport chain (ETC) at complex I, to initiate oxidative phosphorylation. The subsequent flow of electrons through

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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complexes II, III, and IV to the final acceptor O2, promotes pumping of H+ to the intermembrane space at complexes I, III, and IV, to generate a negative mitochondrial membrane potential (ΔΨm) in the matrix. In the coupled respiration, H+ from the intermembrane space are used by complex V (ATP synthase) to generate ATP from ADP and inorganic phosphate. Thus, NADH generation, respiration (oxygen consumption), and ΔΨm are dynamic readouts of mitochondrial metabolism that accurately reflect mitochondrial bioenergetics. A thorough assessment of mitochondrial function is multidimensional, time consuming, and usually requires expensive equipment and specific training. A method to study respiratory complex activities in live cells is currently missing. Individual respiratory complexes have been studied by Western blotting, or by assessing activity in purified complexes, isolated mitochondria or permeabilized cells, out of the multi-regulatory intracellular milieu. However, the amount of each complex does not necessarily correlate with activity, and determinations of substrate utilization during oxidative phosphorylation do not differentiate activities of individual complexes. Measurement of the oxygen consumption rate (OCR) is a method of choice to study mitochondrial respiration and mitochondrial dysfunction [7–9]. In the last decade, the Seahorse technology has been widely adopted to assess basal, maximal, and ATP-linked respiration in adherent cell cultures [10, 11] and isolated mitochondria [12, 13]. Seahorse XF Analyzers are sensitive, robust, medium-throughput, and less labor intensive compared to traditional Clark-type oxygen electrodes [14]. Here, we describe a novel, combined approach to sequentially assess the electron flow through respiratory complexes I–IV in permeabilized and intact cells using Seahorse technology (Fig. 1). The Seahorse analysis is complemented by a highly sensitive and fast method to assess ΔΨm and NADH using plate reader assays (Figs. 2 and 3). Our combined method allows a relatively inexpensive and fast determination of ΔΨm, NADH, and O2 consumption, major readouts of global mitochondrial function in a few hours, using equipment frequently available in laboratories worldwide.

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Materials All solutions are prepared with ultrapure water. Reagents are analytical grade. Unless otherwise indicated, solutions are pH 7.4, filtered and stored at 4  C for 6–8 weeks. The instruments needed are a Seahorse Extracellular Flux (XFe96) analyzer (or similar), and a microplate reader, preferably with spectral scanning capabilities, and CO2/Temperature control (see Note 1).

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A

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100 75 50 25 0

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Fig. 1 Assessment of the activity of individual respiratory complexes in permeabilized and intact cells. Oxygen consumption rates (OCR) in permeabilized cells (top panel) and intact cells (bottom panel), during electron flow experiments as described in Methods. Base: baseline (no chemicals added); A, B, C, and D: ports A, B, C, and D, respectively. Rot: rotenone; Suc: succinate; AA: antimycin A; ASC: ascorbate; TMPD: tetramethyl-pphenylenediamine; Suc-NV: succinate-NV. Averages from three independent experiments 2.1 Buffers for Respirometry 2.1.1 Permeabilized Cells

(a) 3 Mitochondrial Assay Solution with bovine serum albumin (MAS-B): 3 MAS-B contains (mM): KH2PO4 30, sucrose 210, mannitol 660, HEPES 20, EGTA 3, MgCl2 15; and fatty acid-free bovine serum albumin (BSA) 0.9% (w/v). Adjust pH with KOH, filter and sterilize. Store at 4  C (see Note 2). (b) MAS-BP: to 1 MAS-B, add FCCP (4 μM), sodium pyruvate (10 mM), malate (2 mM), and Plasma Membrane Permeabilizer (PMP) (3 nM). Use immediately at 37  C (see Note 3).

2.1.2 Intactg Cells

(a) Solution 1: 10 calcium free electrolyte buffer, contains (mM): MgCl 0.6, KH2PO4 0.5, KCl 5.33, Na2HPO4 · 7H2O 0.5, and NaCl 130; Solution 2: CaCl2 50 (1.8 mM) (see Note 4). (b) Respiratory Medium (RM): add ultrapure water to solutions 1 and 2 to make a 1 solution. Supplement with L-glutamine

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Wavelength (nm) Fig. 2 Spectral scan of TMRM and NADH. Fluorescence spectra of TMRM (top panel) and NADH (bottom panel) in intact cells using a plate reader as described in Methods. Emission wavelengths were 590 nm for TMRM and 440 nm for NADH. Avg  SE from three independent experiments. TMRM: tetramethylrhodamine methyl ester

Fig. 3 NADH and TMRM fluorescence in live cells. Imaging of SNU-449 cells: NADH autofluorescence excited at 720 nm (Multiphoton microscopy; left panel); TMRM (right panel) excited at 561 nm (Single photon microscopy); (scale bar is 10 μm). Note that excitation wavelength for TMRM is very similar to the optimum identified in the spectral scans shown in Fig. 2. NADH wavelength used for multiphoton approximately doubles the optimum identified in the spectral scan because the energy of the electrons in multiphoton is half of single photon (see Note 16)

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(4 mM), glucose (5.5 mM), insulin (100 nM), and fatty acid free BSA (0.3%) (see Note 5). 2.2 Potentiometric Medium (PM)

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Make complete PM by adding TMRM (20 nM) and zosuquidar (1 μM) to HBSS containing (mM): NaCl 137, Na2HPO4 0.35, KCl 5.4, KH2PO4 1, MgSO4 0.81, CaCl2 0.95, glucose 5.5, NaHCO3 25, and HEPES 20. Use the same day of the experiment.

Methods

3.1 Assessment of Respiratory Complex Activity 3.1.1 Permeabilized Cells

(a) Plate cells overnight in a Seahorse XFe96 Cell Culture Microplate (50–60  103 cells per well are optimal for 70–90% confluency monolayers in several cell types). Use whole growth media appropriate for the particular cell line grown at 37  C in 5% of CO2/air. Growth conditions may vary depending on the cell line, but will not affect the assay (see Note 6). (b) Immediately prior to the experiment, prepare 1 MAS-B. Keep it at 37  C. (c) Replace growth media with 175 μl 1 MAS-B incubation medium per well. Place in a 37  C air incubator for 1 h (see Note 7). (d) Prepare MAS-BP. Keep it at 37  C. (e) Setup Seahorse assay protocol for three reads with 0.5 min mixing time, and 3 min measure (see Note 8). (f) Load Seahorse cartridge ports with 25 μl of each ETC inhibitor/substrate (mM) dissolved in MAS-BP: rotenone 0.032, succinic acid 100, antimycin A 0.016, and ascorbate 100 + tetramethyl-p-phenylenediamine (TMPD) 1.0, to ports A, B, C, and D, respectively (see Note 9). (g) Replace incubation medium with MAS-BP, assemble the Seahorse cartridge and start the assay immediately. (h) Normalize results using BCA protein concentration (see Note 10). (i) Analyze results (oxygen consumption rate) using the XFe Wave software (Seahorse Bioscience Inc., MA).

3.1.2 Intact Cells

(a) Same as in Subheading 3.1.1, step a. (b) Immediately prior to the experiment, prepare 1 RM and warm in a 37  C water bath. (c) Replace growth media with 175 μl 1 RM incubation medium per well. Place in a 37  C air incubator for 1 h.

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(d) Setup Seahorse assay protocol to allow three reads, 2 min mix, and 3 min measure per injection. (e) Load Seahorse cartridge ports with 25 μl of each ETC inhibitor/substrate (mM) dissolved in 1 RM: rotenone 0.032, succinate-NV 1.6, antimycin A 0.016, ascorbate 100 + TMPD 1.0, to ports A, B, C and D, respectively (see Note 11). (f) Replace half of the volume of 1X RM incubation medium with fresh 1 RM. After calibration, start the assay. (g) Normalize results by protein concentration or cell number (see Note 10). 3.2 Determination of Mitochondrial Membrane Potential

(a) Plate cells overnight for a 70–90% confluent monolayer in a black wall, glass bottom, 96-well, tissue culture treated plate. Use whole growth media. Typically, 70–80  103 cells per well are optimal for several cell types. Leave 3 wells without cells for tetramethyl rhodamine methyl ester (TMRM) background substraction. (b) Prepare PM. Keep it at 37  C. (c) Remove growth media from 96 well plate gently, washing immediately with PM three times. (d) Incubate cells in PM for 60 min at 37  C in 5% of CO2/air (see Note 12). (e) Perform a spectral scan of TMRM fluorescence using a microplate reader to determine appropriate excitation/emission of TMRM in PM (see Note 13). (f) Replace half of the incubation media with fresh PM and start fluorescence acquisition in the plate reader at excitation/emission 570/590 nm. If optimal wavelengths identified in the spectral scan differ from the 570/590, use those found with the available equipment (see Note 14). (g) Start the assay.

3.3 Determination of NADH Content

(a) Plate cells same as described for mitochondrial membrane potential. Leave wells without cells to make a β-NADH standard curve. (b) Prepare HBSS as described in Subheading 2.2. without TMRM and zosuquidar. Keep it at 37  C. (c) Remove growth media gently. Wash three times with HBSS. (d) Incubate cells in HBSS for 30 min at 37  C in a 5% of CO2/air. (e) Perform a spectral scan of NADH auto-fluorescence using a microplate reader to determine appropriate excitation/emission of NADH in HBSS. Use excitation/emission at 360/430 nm or similar as identified in the spectral scan.

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(f) Dissolve β-NADH in HBSS and make a serial dilution (0–3 mM) for a standard curve (see Note 15). (g) Start the assay.

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Notes 1. The assay can also be done in plate readers without spectral scanning capabilities, temperature, and CO2 control. In that case, it is likely that absolute values will be different, but the assay will still be useful for comparing experimental conditions. Data about basal respiration, maximal respiratory capacity, and ATP-linked respiration can be obtained in separate experiments using a classical respiratory assay using sequential addition of oligomycin, FCCP, and rotenone + antimycin. 2. It is critical to include BSA in the MAS buffer to preserve mitochondrial coupling [15]. Rock BSA-containing tubes gently. Do not vortex to prevent bubbles forming. 3. Prepare MAS-BP 5–10 min before calibration of Seahorse sensor is complete. Minimize the time cells are in MAS-B. MAS based media are not balanced for ionic strength. 4. Store RM buffer stocks parts 1 and 2 separated, since Ca2+ may precipitate during storage. Prepare an aliquot of the working solution the day of the assay. 5. The demands for certain substrates, such as insulin, may differ between cell types. Consider titrating PMP, and measuring cytochrome-C release by western blot for cell lines that may be particularly sensitive to permeabilization. In our setup, up to 3 nM PMP does not permeabilize mitochondrial membranes in HepG2 and SNU-449 cancer cells. 6. Determine the optimal cell density empirically by considering inhibitor/substrate responses for each cell type. If cross cell line comparisons are needed, determine mitochondrial content by quantifying the mitochondria to cell volume fraction, or the average amount of mitochondria per cell. This approach will reduce the possibility of over or underestimations of OCR caused by differences in mitochondrial content. 7. Be careful to aspirate media with a multichannel pipette instead of a vacuum aspirator to prevent cell detachment. All media should be removed by placing the pipette tip against the side of the well without touching the bottom. As described in Note 3, minimize the time cells are in MS-B and the transition to MS-BP. 8. The Mix - Measure cycles can be adjusted. However, the minimum time required to obtain consistent rates is typically 2 min.

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To reduce the probability of cells detaching during the assay (variable for some cell lines), keep the total length of the permeabilized cell assay to less than 1.5 h. 9. TMPD is a Complex IV-specific electron donor. It is important to protect TMPD from light and moisture to avoid oxidation. Make TMPD stock in DMSO before diluting with 1 MAS-BP containing ascorbate. TMPD needs to be maintained in solution with ascorbate to keep its reductive capacity and to prevent autoxidation. An indication that TMPD has been oxidized is a color change in the solution from yellow-green to purple. If possible, prepare fresh working solutions before each assay. Rotenone and antimycin A are inhibitors for complex I and II, respectively. Succinic acid is substrate for complex II and ascorbate and TMPD for complex IV. The final concentration of all compounds in the wells will be diluted to 1 of the port concentrations described in Methods. The higher OCR compared to baseline in the presence of ASC + TMPD can be explained by the low proton motive force (pmf) at complex IV of the ETC. Generally, the lower the pmf the easier it is for electrons to pass along the ETC and reduce O2 to water [16]. 10. To avoid inconsistencies with protein measurements, take into account the BSA content in the MAS-BP media. The permeabilized cell protocol is not compatible with most imaging methods to estimate cell number because cells will be prone to detaching soon after the assay is complete. This is not a concern for the intact cell protocol. Vibrant Green and Hoescht can be used to label nuclei in live cells. 11. Protect Succinate-NV (a cell permeant succinate analog) from light and store stocks at 20  C for up to 3 weeks. Succinic acid is not cell permeant. Both Succinate-NV and Succinic acid are complex II substrates. It is advisable to titrate SuccinateNV for each cell type since high concentrations (typically above 1 mM) inhibit mitochondrial respiration. High Succinate-NV releases formaldehyde which inhibits glycolysis. This factor should be considered if lactate or extracellular acidification rates are determined in parallel. 12. Incubation of cells with TMRM in the low nm range, non-quench mode, typically 10–50 nm, allows a linear association of changes in fluorescence intensity with ΔΨm calculated using the Nernst equation. It is critical to assess the quench limit for each cell type. Intramitochondrial accumulation of TMRM is influenced also by differences in plasma membrane potential (ΔΨp). Ideally, a plasma membrane potentiometric probe should be co-loaded with TMRM to determine the extent to which ΔΨp contributes to changes in TMRM fluorescence [17]. In some cancer cell lines, addition of the

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multidrug resistance pump, zosuquidar, prevents the efflux of TMRM. 13. Fluorophores similar to TMRM can undergo a slight spectral shift when diluted in certain media. An ~3 s spectral scan on a standard microplate reader allows the identification of optimum excitation/emission wavelengths. 14. To reduce the amount of TMRM attached to the walls of culture plates, and to preserve cell attachment, replace half of the initial volume of PM with fresh PM. If plate reader is equipped with temperature and CO2 regulation, wait 15 min after the culture plate is inside the reader to avoid temperature fluctuations that may affect TMRM equilibration. If local plate reader is not equipped with CO2 regulation, exclude sodium bicarbonate from PM. 15. β-NADH fluorescence will respond linearly to the excitation as concentration increases. Culture plates need to be UV-transparent, allowing detection at 340 nm. 16. Determination of TMRM and NADH fluorescence can be confirmed by widefield or confocal microscopy as shown in Fig. 3.

Acknowledgments This work was supported by NIH grant NCI R01CA184456 and SCTR Pilot Project UL1 TR001450-SCTR to E.N.M.; and the Abney Foundation Fellowship from MUSC Hollings Cancer Center, to K.H. References 1. Fang D, Maldonado EN (2018) VDAC regulation a mitochondrial target to stop cell proliferation. Adv Cancer Res 138:41–69 2. Warburg O, Wind F, Negelein E (1927) The metabolism of tumors in the body. J Gen Physiol 8(6):519–530 3. Weinhouse S (1956) On respiratory impairment in cancer cells. Science 124(3215):267–269 4. Ma Y et al (2018) Fatty acid oxidation: an emerging facet of metabolic transformation in cancer. Cancer Lett 435:92–100 5. Ghouri YA, Mian I, Rowe JH (2017) Review of hepatocellular carcinoma: Epidemiology, etiology, and carcinogenesis. J Carcinog 16:1 6. Zhao, Y., Butler, E. & Tan, M. (2013) Targeting cellular metabolism to improve cancer therapeutics. Cell Death Dis 4, e532

7. Brand MD, Nicholls DG (2011) Assessing mitochondrial dysfunction in cells. Biochem J 435(2):297–312 8. Dranka BP, Benavides GA, Diers AR, Giordano S, Zelickson BR, Reily C, Zou L, Chatham JC, Hill BG, Zhang J, Landar A, Darley-Usmar VM (2011) Assessing bioenergetic function in response to oxidative stress by metabolic profiling. Free Radic Biol Med 51(9):1621–1635 9. Rogers GW, Brand MD, Petrosyan S, Ashok D, Elorza AA, Ferrick DA, Murphy AN (2011) High throughput microplate respiratory measurements using minimal quantities of isolated mitochondria. PLoS One 6(7):e21746 10. Connolly NMC, Theurey P, Adam-Vizi V, ˜os JP, Bazan NG, Bernardi P, Bolan Culmsee C, Dawson VL, Deshmukh M, Duchen MR, Du¨ssmann H, Fiskum G,

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Galindo MF, Hardingham GE, Hardwick JM, Jekabsons MB, Jonas EA, Jorda´n J, Lipton SA, Manfredi G, Mattson MP, McLaughlin B, Methner A, Murphy AN, Murphy MP, Nicholls DG, Polster BM, Pozzan T, Rizzuto R, Satru´stegui J, Slack RS, Swanson RA, Swerdlow RH, Will Y, Ying Z, Joselin A, Gioran A, Moreira Pinho C, Watters O, Salvucci M, Llorente-Folch I, Park DS, Bano D, Ankarcrona M, Pizzo P, Prehn JHM (2018) Guidelines on experimental methods to assess mitochondrial dysfunction in cellular models of neurodegenerative diseases. Cell Death Differ 25(3):542–572 11. Heslop KA, Rovini A, Hunt EG, Fang D, Morris ME, Christie CF, Gooz MB, DeHart DN, Dang Y, Lemasters JJ, Maldonado EN (2020) JNK activation and translocation to mitochondria mediates mitochondrial dysfunction and cell death induced by VDAC opening and sorafenib in hepatocarcinoma cells. Biochem Pharmacol 171:113728 12. Salabei JK, Gibb AA, Hill BG (2014) Comprehensive measurement of respiratory activity in permeabilized cells using extracellular flux analysis. Nat Protoc 9(2):421–438 13. Sakamuri SSVP, Sperling JA, Sure VN, Dholakia MH, Peterson NR, Rutkai I, Mahalingam

PS, Satou R, Katakam PVG (2018) Measurement of respiratory function in isolated cardiac mitochondria using seahorse XFe24 analyzer: applications for aging research. Geroscience 40(3):347–356 14. Plitzko B, Loesgen S (2018) Measurement of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) in culture cells for assessment of the energy metabolism. Bio Protoc 8(10):e2850 15. Horan MP, Pichaud N, Ballard JW (2012) Review: quantifying mitochondrial dysfunction in complex diseases of aging. J Gerontol A Biol Sci Med Sci 67(10):1022–1035 16. Berry BJ, Trewin AJ, Amitrano AM, Kim M, Wojtovich AP (2018) Use the protonmotive force: mitochondrial uncoupling and reactive oxygen species. J Mol Biol 430(21):3873–3891 17. Rovini A, Heslop K, Hunt EG, Morris ME, Fang D, Gooz M, Gerencser AA, Maldonado EN (2021) Quantitative analysis of mitochondrial membrane potential heterogeneity in unsynchronized and synchronized cancer cells. FASEB J 35(1):e21148

Chapter 2 Unbiased Millivolts Assay of Mitochondrial Membrane Potential in Intact Cells Chad A. Lerner and Akos A. Gerencser Abstract The mitochondrial membrane potential (ΔψM) is the major component of the bioenergetic driving force responsible for most cellular ATP produced, and it controls a host of biological processes. In intact cells, assay readouts with commonly used fluorescence ΔψM probes are distorted by factors other than ΔψM. Here, we describe a protocol to calculate both ΔψM and plasma membrane potential (ΔψP) in absolute millivolts in intact single cells, or in populations of adherent, cultured cells. Our approach generates unbiased data that allows comparison of ΔψM between cell types with different geometry and ΔψP, and to follow ΔψM in time when ΔψP fluctuates. The experimental paradigm results in fluorescence microscopy time courses using a pair of cationic and anionic probes with internal calibration points that are subsequently computationally converted to millivolts on an absolute scale. The assay is compatible with wide field, confocal or two-photon microscopy. The method given here is optimized for a multiplexed, partial 96-well microplate format to record ΔψP and ΔψM responses for three consecutive treatment additions. Key words Mitochondrial membrane potential, Plasma membrane potential, Mitochondrial biogenesis, Fluorescence microscopy, Live cell microscopy, Cell culture, Tetramethylrhodamine methyl ester, TMRM, Bis-oxonol, MitoTracker, Cellular heterogeneity, Single cell

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Introduction The mitochondrial membrane potential (ΔψM) is a commonly assayed bioenergetic variable, but typical fluorescence assays in intact cells are prone to data misinterpretation due to factors other than ΔψM contributing to the readout. ΔψM is the main component of the proton motive force across the mitochondrial inner membrane that drives mitochondrial ATP synthesis [1]. ΔψM directly regulates or powers mitochondrial quality control [2], protein import [3], transport of certain solutes [4], and is linked to retrograde signaling by mitochondrial peptides [5]. Therefore,

Supplementary Information The online version contains supplementary material available at [https://doi.org/ 10.1007/978-1-0716-2309-1_2]. Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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the magnitude of ΔψM is informative of bioenergetic function, mitochondrial fitness, or damage. Fluorescence probes TMRM, TMRE, rhodamine 123, and JC-1 are broadly used to assay ΔψM in in vitro or ex vivo intact cell specimens [6–9], such as cell and tissue cultures, isolates [10], and biopsies [11]. When fluorescence intensities are directly analyzed, these methods are semi-quantitative [6], meaning that the scope of data interpretation is limited, e.g. to “sudden increase” or “sudden decrease” of ΔψM when using quench mode probes (see Note 1), or comparisons are possible only with strict assumptions, e.g. same cell size, geometry, and constant ΔψP when working in non-quench mode [12]. The rhodamine probes follow well-understood laws of chemiosmosis, and this is the basis of the fully quantitative approach described below [12]. In contrast JC-1, although attractive due to its ratiometric fluorescence, is unsuitable for unbiased ΔψM determination [6], because of unpredictable (or not yet understood) principles that control its heterogeneous phase aggregation. Semi-quantitative use of another class of mitochondrial labels, the MitoTrackers have been also reported, despite the mitochondrial accumulation of these labels being only partially ΔψM-dependent and exhibit fluorescence quenching [13] and retention in mitochondria after depolarization [14]. Other alternatives, such as radioisotope labeling of bulk, suspension cultures, or quantitation of quench mode signals were discussed previously [12]. We have described these constraints and pitfalls of alternative methods in detail in [12, 15]. The method presented here provides unbiased, absolute calibrated (millivolts) values of both ΔψP and ΔψM in intact, cultured or immobilized isolated live cells [12, 15]. The key advantage compared to semi-quantitative techniques is that ΔψM is determined in an absolute scale with no a priori assumptions on cell geometry or ΔψP, allowing comparisons of specimens, using micro-scale samples, down to single cells. The technology builds on and extends earlier quantitative radioisotope, electrochemical and fluorescence ΔψP and ΔψM determinations (see [12]). The approach is fluorescence microscopic, microplate-compatible, uses a pair of common fluorescence probes, and computational image and data analysis. These probes are tetramethylrhodamine methyl ester (TMRM) and FLIPR Membrane Potential Assay Explorer Kit (from Molecular Devices, also referred to as plasma membrane potential indicator, PMPI [12, 16]; see Notes 2 and 3). The method describes a time course recording paradigm in adherent or immobilized cells that establish internal calibration points at the end of the recording, allowing conversion of preceding fluorescence intensities to millivolts. Then, this conversion is performed by computational image and data analysis using interactive software modules designed for this purpose in Image Analyst MKII (Image Analyst Software). Furthermore, a calculation spreadsheet provided with the software allows planning for three consecutive drug additions into each assay well. Fluorescence time lapse recording may be

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performed using wide field, confocal or two-photon microscopy, and manual liquid handling to expose cells to drug additions and calibration media (see Note 4 on microplate readers and flow cytometry). This method has been proven to work robustly on different specimens, including pancreatic β-cell primary cultures [15, 17, 18] and cell line [19], embryonic and neural stem cells [20], osteoblasts [21] and hepatocellular carcinoma and lung adenocarcinoma cells [22], and also on a variety of microscope systems.

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Equipment 1. Inverted fluorescence microscope with 10
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fluorescence-optimized, high NA (0.5–0.75) lens (see Note 5), high-sensitivity cooled CCD or sCMOS camera, softwareswitched LED light source or other white light source with filter wheel and fast electronic shutter (see Note 6), real time autofocus (see Note 7), x,y-motorization with microplate adapter, heated environment chamber (see Note 8). Ideal fluorescence filters for the FLIPR ΔψP probe and TMRM, respectively, are: for excitation 509/22 nm (bandpass filter, given as center/bandwidth, e.g. from Semrock) and 586/20 nm (we use these in an 18 incident angle in a Sutter Lambda 821 LED light source with 506 nm and 561 nm LEDs) or 500/24 nm and 580/14 nm using a Xe-arc light source; dichroic: 459/526/596 triple edge (Semrock) for both probes, or separate 520 and 593 nm high pass dichroic mirrors; emission: 542/27 nm and 641/73 nm. Alternatively, a multiband emission filter can be built by the combination of an FF01-475/543/702 and a BLP01-514R filter, allowing fast, solid-state switching between the two channels using the LED light source. As a basic alternative, a standard FITC or YFP filter cube for FLIPR, and a Cy3 or Texas Red filter set for TMRM may be used at the cost of a greater spectral crossbleed. For setting up the sample under eyepiece, a red filter, e.g. 600 nm long pass is dropped into the bright field illumination pathway to protect specimen from photodamage. 2. Alternative: Biotek Citation 5 plate reader microscope with standard FITC (or YFP) and Cy3 LED-based illumination, laser autofocus, and environment control. 3. Alternative: two-photon microscope, 20
high NA water immersion lens (for upright configuration) 910–930 nm mode-locked excitation, non-descanned detectors with a 593 nm beamsplitter to detect two emission channels simultaneously using emission filters as specified in #1 (see Note 9). 4. An inverted laser scanning confocal microscope is required for measurement of mitochondria:cell volume fraction and

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apparent activity coefficient ratio (see below), and it is also an alternative equipment for the ΔψM. Equipped with 488 nm, and 561 nm laser lines, and detection at 521–558 nm and 597–668 nm [22]. Alternatively, 543 nm excitation may be used for TMRM at adjusted detection bands, at a greater spectral crossbleed. If using a tunable white laser light source, set up light path as given above “ideal” for wide field microscopy. Real-time autofocus and an environment chamber are critical for stable recordings. Perform ΔψM recordings with an open pinhole. 5. 8- or 12-channel handheld pipettor (200 μL) that fits between the microscope stage and the ceiling of the environment chamber. 6. Motor pipettor for serological pipettes. 7. Ambient air 37  C incubator, ideally with a rocker. 8. 37  C waterbath. 9. For cell immobilization a swing-bucket refrigerated centrifuge with microplate adaptors. 10. Software: Image Analyst MKII (Image Analyst Software, Novato, CA), Microsoft Excel and optionally GraphPad Prism.

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1. Tetramethylrhodamine methyl ester (TMRM), dissolve in DMSO at 5 mM and then dilute to prepare 50 μM stocks. Store 100 μL aliquots at 20  C (see Notes 3 and 10). 2. FLIPR Membrane Potential Assay Explorer Kit (#R8042, Molecular Devices), dissolve one vial in 10 mL H2O. Store at 20  C, aliquoted for one time use size (see Notes 3 and 9). 3. MitoTracker Red CMXRos (50 μg vials), prepare 100 μM stock in DMSO. Store 100 μL aliquots at 20  C. 4. Calcein AM (50 μg vials), prepare 1 mM stock in DMSO and store at 20  C.

3.2 Assay Media Components

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potentiometric medium (2
PM): Prepare 1 L of 7 mM KCl, 2 mM MgCl2, 0.8 mM KH2PO4, 40 mM N-[Tris (hydroxymethyl)methyl]-2-aminoethanesulfonic acid (TES), 1 mM NaHCO3, 2.4 Na2SO4, pH 7.4 set by NaOH at 37  C (see Notes 8 and 11). Sterile filter (see Note 12) and store at 4  C. Alternatively use culture-medium-formulated 2
PM. This may be done by omitting the following components of any solid media formulation: NaCl, CaCl2, phenol red, folic acid, riboflavin). Then, the 2
PM is made by dissolving double amount of solid media, and TES, NaHCO3 and

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CaCl2 as above specified. Add NaCl, so the total Na+ in this medium is 120 mM less than in the original formulation, when calculated for final dilution. Set pH and store as above. 2. 2
NaCl and 2
KCl, prepare 1 L and 250 mL 240 mM solutions in H2O, sterile filter and store at RT (see Note 12). 3. CaCl2 and D-glucose, prepare 10 mL of 1 M solutions in H2O, sterile filter, and store at RT (see Note 12). 4. Sodium tetraphenylborate, (TPB), prepare 1 mM stock in H2O. Store 100 μL aliquots at 20  C. 5. Zosuquidar, optional to inhibit glycoprotein-P pumping TMRM, prepare 25 mM stock in DMSO. Store 50 μL aliquots at 20  C. 3.3 Calibrant Components

1. Mitochondrial depolarization cocktail (MDC; see below at Note 28) stock components: first, prepare small amounts (see Note 13) of the following stock solutions separately in DMSO (alternatively EtOH may be used): 10 mM valinomycin, 10 mg/mL oligomycin A, 20 mM antimycin A (or myxothiazol), 2 mM carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP), and optional components: 500 mM IAA-94, 100 mM DIOA, and 200 mM bumetanide. Then make MDC by mixing these components based on Table 1. Store at 20  C (see Note 10). 2. KCl, prepare 10 mL of 2 M solution in H2O, sterile filter, and store at RT (see Note 12). 3. Paraformaldehyde solution (8% in H2O): (a) Use 8 g paraformaldehyde for 100 mL distilled water. (b) Heat to 60  C while stirring. Add drops of 10 M NaOH until the solution is transparent. (c) Cool to room temperature. (d) Filter into aliquot tubes. Store frozen at 20  C. (e) Thaw before use in 65  C water bath. (f) Adjust to pH 7.4. (g) Discard after 1 week of storage at 4  C. 4. Gramicidin, prepare 100 μL of 10 mg/mL stock solution in EtOH. Store at 20  C.

3.4 Cell Culturing Reagents (Optional)

1. Matrigel or Geltrex, aliquoted for one time use size at 0  C, store at 80  C (see Note 17). 2. Polyethyleneimine dissolved in H2O (PEI; 1:15000 w/v %; Sigma-Aldrich #P3143). 3. PBS.

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Chad A. Lerner and Akos A. Gerencser

Table 1 Mitochondrial Depolarization Cocktail (MDC; 1:1000) stock Component

Final (μM)

Stock (mM)

Volume (μL)

Core MDC components

Valinomycin Oligomycin FCCP Antimycin

1 2 μg/mL 0.1 2

10 10 mg/mL 2 20

10 20 5 10

Anti-swelling components (optional)

IAA-94 DIOA Bumetanide

100 10 50

500 200 200

20 5 25

Add DMSO (for 100 μL total volume)

3.5

Consumables

5

1. Deep-well reservoir (pyramid bottom, square shape, 2 mL, VWR# 75870-792, plus matching lids, such as Corning #3931). 2. Glass inserts for reservoir (0.7 mL, Chemglass# CV-13500008, VWR# 75820-668). 3. Glass vials 60 mL and 8 mL (see Note 14). 4. Coverglass-bottomed 96-well microplate. Volumes are calculated for the Corning half-area 96-well high content imaging plate (#4580). Alternative: Greiner SensoPlate 96-well glass bottom microplate (see Note 15). 5. Teflon tape (1 mil thick, 200 wide roll, such as CS Hyde Company #23-1FEP-2-50). 6. Can of cleaning duster. 7. Ice pack bricks (warm bricks hereafter), kept at 37  C in the ambient air incubator. 8. Trough reservoir (25 mL) for multichannel pipetting. 9. 50 mL polypropylene conicals, various sizes of polypropylene (Eppendorf) tubes. 25 ml serological pipettes. 10. 25 ml serological pipettes.

4

Methods The following method applies to adherent cells grown to sub-confluent densities (see Notes 16 and 17) or freshly isolated or suspension cultured cells immobilized in a 96-well microplate. We recommend that the assay layout comprises of 1–4 columns of six wells (avoiding edges in a 96-well microplate) on the side of the handedness of the experimenter, or where there is a sufficient space available for pipetting within the environment chamber of the microscope. The well labeled “A1” below refers to the top left of

Millivolts Mitochondrial Membrane Potential in Intact Cells

17

this range. The method is given to obtain a time course of potentials comprising of four segments (of desired duration, e.g. ~30 min each); a baseline, and subsequently (optionally) up to three consecutive test compound additions and respective recordings. This is followed by a sequence of calibrant additions. The calibration steps are required in every recording in order to calculate millivolt potentials. To avoid cell wash-off while providing adequate mixing, most additions are performed by removing half of the assay volume in each well and adding fresh PM with treatment of choice at 2
the concentration desired. For consecutive additions, an Excel worksheet-based dilutions table is described to calculate carry over of previous treatments into each new half-volume replacement, thereby preserving concentrations of previously added drugs as further additions are made. The mixing paradigm below also ensures that the probe concentrations remain constant during the experiment. To calculate millivolts from fluorescence intensities, a minimum of two calibration points are required at two differing potentials. However, in intact cells only the zero potential can be precisely established. In this sense, the calibration paradigm is a workaround for establishing a non-zero potential calibration point. We have developed multiple ΔψM calibration paradigms that differ in the actual calibration points, either to simplify the original method or to overcome specimen-related obstacles. Here, a complete calibration method is presented that provides millivolt calibration for both ΔψP and ΔψM without the knowledge of starting, baseline potentials. The alternative methods are discussed below in Notes 42–46. Furthermore, there are two parameters specific for the specimen, which are required for all calibration paradigms, but cannot be determined from the time course recording. These are the mitochondria:cell volume fraction (VF) and the apparent activity coefficient ratio (aR’; see definitions in [12]). The former is a stereologic term, while the latter collectively describes a matrix to cytosol ratio of chemical activities of TMRM, its binding to mitochondrial membranes and the dilution of fluorescence by ultrastructural details (e.g. crista density). These parameters are measured by confocal microscopy in Tasks #13–14. Time required to perform a 3-addition, multiplexed experiment is ~8 h, not including preceding media and stock preparations, following image and data analysis, and accessory determinations (VF, aR’, spectral crossbleed). The protocol is organized into tasks below, illustrated by the timeline in Fig. 1. 4.1 Task #1: Pilot Experiments

In light of the protocol below, consider these pilot experiments: 1. Measure the volume of the assay well when it is filled flush to the rim.

18

Chad A. Lerner and Akos A. Gerencser

Prepare stocks Design layout (dilutions template) Cell culturing matching the template or plate coating for immobilization Prepare assay media Wash cultured cells or immobilize fresh cells Prepare additions reservoir Assemble and warm up final wash reservoir Optional immunocytochemistry and re-imaging

Final wash

Continue incubation on the microscope, set up recording Start baseline recording additions Task 1,2

3

4 or 5

Cells are cultured

6

7.1

7

8.1-4

Cells are preincubated in PM

8.5

calibration

8.6-10

8.6.11-22

data analysis 9-11

Cells are on the microscope in PM

Critical timing:

5m -3 in 0m in

0

30min

2h

2h-4h next day

-4

-3h

0m

-4h

-9

previous day

in

Image acquisition

There is sufficient time to complete this step.

Sample and additions reservoir are warm on the microscope. Calibrants are warm on the microscope.

Fig. 1 Timeline of the assay

2. Test the cell attachment against the wash paradigms (e.g. Task #4). 3. Optimize probe concentrations and microscopy settings for minimal necessary illumination intensities and maximal acquisition speed (see Notes 5, 18, and 19). 4. Determine how to save image data so it is compatible with the analysis in Task #9 (see Note 20). 5. Determine the cycle time of microscopy in partial microplates (see Note 31). 6. Determine spectral crossbleed between FLIPR and TMRM (see Task #12). Critical: Do not change optical configurations after measuring crossbleed (see Note 52). 7. First experiments may use a single well or single column of wells and only a baseline recording followed by the calibration sequence. 4.2 Task #2: Additions Design

Skip this section if no test compound additions are planned, i.e. only baseline and following calibration are recorded. Design additions and treatment dilutions in advance, and match it to the cell culture plating pattern using a 96-well microplate. In this task, dilutions and volumes of test compound stocks are calculated in order to prepare 2
concentrated additions with carryover of previously added treatments (see Note 21). These calculations are used in Task #6, and the dilutions paradigm is illustrated in Fig. 2b.

Millivolts Mitochondrial Membrane Potential in Intact Cells

A

2×PM Table 2 #1 +D-glucose +TMRM +FLIPR +tetraphenylborate +zosuquidar +CaCl2

B Addition #1

PMK Table 2 #2

2×NaCl 30 ml

PMPFA Table 2 #3

PM Table 2 #4

Volumes used per assay well 80ul ~2.5ml

145ul

C

+3× MDC stock 3 ml

+PFA +KCl 2M +gramicidin 2 ml

2 ml 2 ml

+1× MDC stock

Addition #3

Addition #2 50μl 100μl

350μl

30 ml

2×KCl

19

2x

4x

8x

700μl

350μl

50μl

2x

2x

300μl

600μl

2x 200μl

MDC Table 2 #5 50ul

discard 50μl add 50μl and mix

PBS

100μl

50μl

100μl

Fig. 2 Media preparation and liquid handling paradigm for keeping constant probe concentrations. (a) Flow chart representation of supplementation and dilution of 2
PM resulting in the assay medium PM and the calibrants MDC, PMK and PMPFA in Task #3 (see Table 2 and Notes 26–28). (b) Scheme of preparation of three consecutive 2
concentrated compound additions with carryover of previously added compounds in Task #6 (see Note 21). The vessels depict reservoir glass inserts. Top row: First, fill inserts with the indicated volume of PM with addition compounds at the indicated times of final concertation. Then transfer volumes indicated by the arrows on the top, resulting the bottom row. (c) Illustration of liquid handling paradigm during recording in Task #8.7. Three wells of a glass-bottomed microplate are depicted with the edge well filled with PBS. Additions prepared in B are mixed into the assay volume by half replacement

1. Using Image Analyst MKII main menu “Tools/Membrane Potential Assay Dilutions Worksheets” open a dilutions template and save it as a new Excel file (“dilutions template.xlsx”). Using this workbook: 2. Edit “Plate” tab only above row 31 to enter treatments for each well. Critical: do not use the cut command to modify the table, only copy/paste. Do not remove or add rows. 3. Columns A,G,M: well labels are arbitrary. If matching image metadata are available, these are used for automated well to condition assignment below (Task #9.4).

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Chad A. Lerner and Akos A. Gerencser

Table 2 Preparation of potentiometric media and calibrants. See Fig. 2A Medium

Component

Final (in 1
)

Stock

Volume

#1 Supplemented 2
PM In 50 mL conical

2
PM D-glucose TMRM FLIPR Tetraphenylborate, Na Zosuquidar (optional) CaCl2

1
5.6 mM 7.5 nM 1:100 1 μM 1 μM 1.8 mM

2
1M 50 μM 100
1 mM 25 mM 1M

40 mL 448 μL 12 μL 800 μL 80 μL 3.2 μL 144 μL

#2 PMK In 8 mL glass vial

Supplemented 2
PM 2
KCl MDC stock

1
120 mM 1:1000

2
240 mM 1000


2 mL 2 mL 4 μL

#3 PMPFA In 8 mL glass vial

Supplemented 2
PM PFA solution 2 M KCl Gramicidin

1
~2% 120 mM 5 μg/mL

2
8% 2M 10 mg/mL

2 mL 1880 μL 120 μL 2 μL

#4 PM In 60 mL glass vial

Supplemented 2
PM 2
NaCl

1
120 mM

2
240 mM

30 mL 30 mL

#5 MDC In 8 mL glass vial

PM MDC stock

3:1000

1000


3 mL 9 μL

4. Columns B-E, H-K, N-Q: enter compound name, desired final concentration, stock concentration, and unit. Final and stock concentrations are in the same units here. 5. Leave unnecessary rows or addition groups empty. 6. Revise assay and reservoir volumes in rows 27–29, values in red, if needed. 7. Tables from row 31 are automatically populated. In the table on the left (from cell A31) assay wells are shown in pairs of rows, and the three consecutive additions in pairs of columns. For each well/addition, compound name, stock concertation, dilution factors, and volumes are indicated on the top, and the volume of PM on the bottom (see Note 21). 8. Alternatively, one may use the same addition in multiple wells or may prefer to pipette larger volumes than calculated. To this end, a tally is automatically created on the right (from cell J30). The last two columns of the tally contain practical volumes of compound stock and PM to mix. 9. The “Checklist” worksheet organizes the compound additions in a table to aid organization of the additions reservoir plate below.

Millivolts Mitochondrial Membrane Potential in Intact Cells

21

10. The “Condition Names” worksheet matches image metadata and descriptive condition names generated from the compound names and concentrations when a column of well labels (see step #3) is pasted into A27. 4.3 Task #3: Potentiometric Media and Calibrants

The preparation of the potentiometric medium (PM) and the calibrants (MDC, PMK and PMPFA) is the first task on the day of the assay. Prepare fresh assay media and calibrants for each experimental day (see Note 22) using the stocks defined in Section #3. This task is illustrated in Fig. 2a. 1. In a 50 mL polypropylene conical tube supplement 40 mL 2
PM according to Table 2 section 1, and mix with vortexing (see Notes 8, 12, 23, and 25). 2. Take 2 of the 8 mL glass vials, and transfer 2 mL of supplemented 2
PM into each and label as “PMK” and “PMPFA.” These will be later completed as calibrants (see Note 26 and 27). Critical: PM and all calibrants are kept in glass vials to prevent adsorption of TMRM to the side walls. The 2
PM is kept only shortly in the polypropylene tube. 3. Take a 60 mL glass vial and using the same serological pipette, first add 30 mL of 2
NaCl, then 30 mL of the supplemented 2
PM, label as “PM” (now 1
) and mix it with vortexing. Critical: For each new batch of 2
PM verify that the pH is 7.4 at 37  C after making PM. If necessary, an amount of NaOH or HCl that corrects the pH can be included in step #1. 4. Transfer 3 mL of PM into a third 8 mL glass vial and label it as “MDC” (see Note 28). 5. Add 2 mL of 2
KCl to the to the PMK tube. 6. Store calibrant vials and remaining supplemented 2
PM in a dark drawer until use. Wrap the PM vial in aluminum foil and place it in a 37  C waterbath. 7. Thaw PFA solution, unless available from previous days. 8. Now, or before use, complete supplementation of MDC, PMK, and PMPFA based on Table 2.

4.4 Task #4 Assay Plate of Cultured Adherent Cells

Prerequisites for preparation of the assay plate of cultured adherent cells are a sub-confluent (see Note 16) density culture that has been grown in an appropriate (coated) coverglass- or plastic-bottomed microplate (see Notes 15 and 17), and cultures were plated in a well pattern matching the experimental design in Task #2. The culture had a long-enough growth period for strong attachment, but there are occasional gaps in the monolayer, that will be used for background subtraction. The wells immediately outside of the assayed range (e.g. edges) are filled with PBS. Critical: avoid touching and

22

Chad A. Lerner and Akos A. Gerencser

keep the bottom of the microplate spotless during culturing and assaying. 1. Place two 37  C warm bricks, and a waste container on the bench or into a TC hood (see Note 29). Critical: To protect live cells photosensitized by TMRM, turn off any over the bench or TC hood light and avoid direct sunlight. Ambient lighting is OK, do not work in darkness. 2. Place a trough reservoir on the warm brick and decant ~15 mL of warm PM (from Task #3.6). 3. Place the culture plate on the other warm brick. 4. Set the multichannel pipette to 200 μL, and remove all culturing media from the first assay column. Critical: pipette out of the corner of the well and rock the pipettor slightly so all growth media are removed in all wells. Small remaining amounts may interfere with the assay (see Note 25). Critical: do not use vacuum aspirator unless the culture is very robust. 5. With changed tips add 200 μL of PM, wash the side walls and fill the wells flush to the top. This is 205 μL when using the half-area 96-well microplate (Corning #4580). Critical: Do not contaminate PM with growth media (see Note 25). 6. Repeat steps #4–5 for all columns. 7. Do two more washes by repeating steps #4–6 but with setting the pipettor to 180 μL. Critical: do not remove all medium during washes other than the first wash, unless the culture is very robust. 8. Remove 50 μL from each well (to avoid the PM touching the lid), cover microplate with lid and place into a 37  C ambient air incubator on a rocker (see Note 8). Critical: cell and all reservoir plates need to be covered by lid during all incubations to prevent evaporation. 9. Incubate 1–4 h. 10. During incubation time perform the following Task #6 and Task #7.1. 4.5 Task #5 Alternative: Assay Plate of Immobilized Cells

Preparation of the assay plate using suspension cultured or freshly isolated cells requires a live cell suspension washed with PBS or kept in isolation medium, on ice. Previous day: 1. Coat wells of a coverglass-bottomed microplate overnight by incubating with 50 μL/well 1:15000 (w/v) PEI solution in a TC-incubator. Previous Day: 2. Wash PEI-coated wells 2
with 150 μL PBS.

Millivolts Mitochondrial Membrane Potential in Intact Cells

23

3. Remove all liquid with aspirator and store the coated plate at 4  C until use. Critical: The plate will need to be cold when cells are added. 4. Thaw an aliquot of Geltrex on ice. Critical: Geltrex or instruments handling it should be ice cold at all times, with the first time Geltrex being exposed to higher temperatures in step #13. 5. Put 10 mL PM in a 50 mL conical on ice. 6. Put 20 mL PBS in a 50 mL conical or small beaker on ice. Use this to cool pipette tips for handling Geltrex and cells. 7. Pellet cells using cell culturing protocols and resuspend them in 1 mL ice-cold PM (see Note 25). 8. Place the coated glass-bottomed microplate on a 4  C cooled “ice” pack. 9. Dilute cells to a density to plate each well in 45 μL. For the halfarea plate, 20,000 cells per well is a good starting point. Thus, dilute the suspension to 4.4
105 cells/mL density. 10. Cool a pipette tip in the ice-cold PBS and add 10% Geltrex to the diluted cell suspension. 11. Using a cooled pipette tip, add 50 μL cell suspension to the range of assay wells. 12. Spin the plate for 5 min at 500
g in a 4  C refrigerated swingbucket centrifuge. 13. Incubate 1–4 h in a 37  C ambient air incubator (see Note 30). 4.6 Task #6: Preparation of Treatment Additions

Skip this section if no treatment additions are planned, except for performing step #2. In this task, 2
concentrated additions with carryover of previously added treatments are prepared using a dilutions paradigm illustrated in Fig. 2b and calculations from Task #2. Critical: This is a time-consuming task, depending on the number of wells and additions it may take 1–2 h (Fig. 1). Staggering with assay plate washes and preincubations is recommended. Critical: The additions reservoir plate prepared here must be ready and warmed up to 37  C by the time the baseline recording ends (see below). 1. In a 96-well deep-well reservoir microplate, place 0.7 mL glass inserts in the same pattern of the assay wells and repeat this in groups for each addition. This should also match the “Checklist” layout in Task #2.9. 2. Add two more columns of inserts for calibrants on the right (or may use a second reservoir plate). These will remain empty for now. 3. First work on addition #1 below, filling the left-most group of inserts.

24

Chad A. Lerner and Akos A. Gerencser

4. There are two ways of filling the inserts. Either pipette volumes calculated in the “Plate” worksheet (see Task #2) from cell A31 directly into the inserts or use the tally table from cell J30. For the latter, using an appropriately sized tube, pipette the volume of compound and PM indicated in columns P and Q, respectively. Then dispense the volumes indicated in row 31 into reservoir inserts. Critical: Do this insert by insert, and do not leave PM in a polypropylene tube for a prolonged period of time (see Note 24). 5. When all inserts are filled, mix their content using a 200 μL multichannel pipette using changed tips between columns. 6. If doing consecutive additions, using the multichannel pipette with changed tips, transfer volumes calculated in row 83 from each of the addition #1 wells to the corresponding addition #2 and #3 wells (e.g. from A1 of addition #1 to A1 of addition #2). By default, this is transferring 350 μL from additions #1 to #2 and 50 μL from additions #1 to #3 (see Fig. 2b). 7. If doing consecutive additions, with new tips, mix addition #2 and transfer volumes calculated in row 84 from each of the addition #2 wells to the corresponding addition #3 wells. The default volume is 100 μL (see Fig. 2b). 8. Optionally, the two last columns (from step #2) can be filled with fully supplemented MDC and PMK (Table 2) now, by distributing all made calibrant in the six reservoir inserts of each column. Alternatively, do this at the latest in Task #8.11 below. Critical: Have sufficient volume of calibrant for all columns of wells in the experiment. 9. Cover the reservoir microplate with a lid and store in a dark drawer until use. Place it in the microscope’s environment chamber latest before starting baseline acquisition. 4.7 Task#7: Final Wash of the Assay Plate

1. Approximately 90 min before the planned start of recording, assemble another 96-well deep-well reservoir for a “final wash” of cell cultures using PM. Place 0.7 mL glass inserts in the pattern of the assay wells once, fill them to capacity with PM, and warm it to 37  C in the ambient air incubator, covered by a lid (see Note 30). 2. About 45 min before the planned start of the recording, place this reservoir and the preincubated assay plate each on a warm brick. 3. Fill assay wells flush with PM. This is 205 μL when using the half-area 96-well microplate (Corning #4580). If using larger wells, fill flush and then set to 200 μL by removing the difference.

Millivolts Mitochondrial Membrane Potential in Intact Cells

25

4. Wash wells 2
with 180 μL PM without contaminating the reservoir inserts. 5. Fill assay wells flush with PM again, and set well volume to 100 μL by removing the difference from the known full-well volume. This will be the starting volume of the assay. 6. Transfer the assay plate into the warmed-up stage and environment chamber of the microscope (see Note 29). With a brief burst of duster can, remove lint or dust from the bottom of the plate. 7. Replace microplate lid with a small piece of Teflon film, that can be lifted or slid off wells for additions. A duster can may be used to remove any debris from the film. 8. Place the lidded additions reservoir from Task #6.9 into the environment chamber. 9. Allow for at least 30 min of thermal equilibration time before starting baseline recording. 4.8 Task #8: Potentiometric Recording

Prerequisites for performing the fluorescence microscopic time lapse recording are that the microscope has been already configured for recording two-channel, TMRM and FLIPR (in this order) time courses with attenuated illumination (see Notes 5 and 19), in multiple x,y-positions, and single z-plane with auto-focusing (see Notes 7 and 5). The assay layout was designed with the cycle time of the time course in mind (see Note 31). The format of image data to be saved supports time stamps that recreates a correct time axis when analyzed in Task #9 (see Note 20). 1. Find cells under eyepiece and brightfield illumination and coarse focus. Critical: prolonged brightfield illumination will depolarize ΔψM in TMRM-loaded cells. Accommodate eyes to darkness and attenuate intensity. Ideally, use a red filter in the bright field illumination pathway. 2. In an area that is not going to be imaged, fine focus cells using snapshots of TMRM, and set up the real-time auto-focusing. Critical: Attenuate illumination intensity, and do not use continuous fluorescence excitation to examine cells. 3. Critical: Adjust illumination intensity and exposure time (or scan speed) to levels not causing photo damage during recording (see Note 19 and constraints due to spectral unmixing in Note 52). Critical: Do not saturate detector. 4. Set up x,y-positions for the part microplate recording. Use a grid of coordinates, or manually find appropriate view fields using attenuated bright field illumination. Multiple view fields per well are recommended without overlap of the illuminated areas. Well naming such as “A1_1” and “A1_2” is compatible




Complete
(known k)

Baseline & Zero

(
) Baseline & MDC or CDC Complete
Complete
(known k)

Complete iterative Complete
Complete
(known k)

Complete




Complete with known kP (Ksteps)

Baseline ΔψP Baseline ΔψP, kT




kT

kT, (PN  0)

(PN  0)

kT















Baseline ΔψP and ΔψM, kT










Assessing cell-to-cell heterogeneity of ΔψM assuming ΔψP (and kT) are the same

No cycle time constrain, more positions are possible

Fewest assumptions and constraints

More robust than “. . .Goldman” in some cell types + long cycle time, ΔψM depolarized by additions

For generic use + long cycle time, ΔψM depolarized by additions

Previously measured (assumed) parameters Indication/rationale


















PMK PMPFA TMRM Response decay to K+steps




MDC





ΔψM

ΔψP

Time course characteristics

Complete
Complete with
known kP (K- Complete (known k) steps) Goldman using Neural Network

Calibrants required

Calibration method

Table 3 Comparison of calibration paradigms

26 Chad A. Lerner and Akos A. Gerencser

Millivolts Mitochondrial Membrane Potential in Intact Cells

27

with the automated pooling of technical replicates below (see Tasks #2.3 and #9.4). 5. Verify focus and culture integrity in each position with a few snapshots of TMRM or FLIPR fluorescence (the latter is typically dim at the baseline). 6. If both assay and reservoir plates are thermally equilibrated in the environment chamber, start baseline recording (see Note 32). Set the acquisition software for a ~ 30 min recording (see Notes 33 and 34). 7. When the recording is complete, working column-by-column, use the multichannel pipettor to remove 50 μL PM (or the volume according to row 28 of the “Plate” worksheet), and using the same (now warmed up) tip, replace PM from the corresponding column of the additions reservoir, and mix with five gentle strokes (see Fig. 2c). 8. Repeat step #7 for each column using fresh pipette tips. Critical: place Teflon film and reservoir lid back at the end of the addition to prevent evaporation. 9. Record next segment of the time course (see Notes 33 and 34). 10. Repeat steps #7–9 for all additions. 11. Fill up the calibrant reservoir inserts (see Task #6.2) with warmed-up supplemented MDC and PMK calibrants if this have not been done earlier. One column of inserts is filled with MDC and the second column with PMK. These calibrants must be at 37  C when the last segment (or the baseline of there are no additions) is completed. Do not fill PMPFA into inserts at this point, but place the PMPFA vial into the environment chamber to warm it up. 12. Add 50 μL MDC without any PM removal, by gently ejecting it toward the bottom of the well and quickly but gently mixing by 5 strokes (see Note 28). 13. Start another 30 min recording. Critical: Start recording promptly, not to miss the decay of TMRM fluorescence intensity, unless alternative calibration is used (see below in Table 3 and Note 43). 14. If alternative calibration is used with no K+-steps (see below in Table 3 and Notes 42 and 46), fill empty calibrant inserts with PMPFA and jump to step #22. 15. Add 20 μL PMK into each assay well and mix. 16. Wait 1 min and record 3 time points. 17. For each assay well, remove 40 μL medium, add 40 μL PMK and mix. 18. Wait 1 min and record 3 time points.

28

Chad A. Lerner and Akos A. Gerencser

19. For each assay well, remove 75 μL medium, add 75 μL PMK and mix. 20. Wait 1 min and record 3 time points. 21. Discard remaining PMK in the additions reservoir and fill the same inserts (no washing) with warm PMPFA. 22. For each assay well, remove 90 μL (70 μL if no PMK addition) medium and add 80 μL PMPFA (added at 1:1 mixing to the contents of the well; see Note 27). 23. Record for 30 min or longer, until a plateau in FLIPR fluorescence intensity is observed. This is the last segment of the experiment (see Note 35). 4.9 Task #9: Image Analysis

1. Open the entire recording in Image Analyst MKII (see Note 20). If the time course was recorded in separate files, use multifile select. This results in a “Multi-Dimensional Open” dialog. 2. In the “Multi-Dimensional Open” dialog use the drop-down box in the top left to select any x,y-position and press “Open” to inspect the recording for integrity. The resulting pair of “Image Windows” show TMRM and FLIPR fluorescence. Critical: Ensure that when pressing “Open” a time series from a single position is opened, e.g. by check marking “Merge in Time” unchecking “Positions as Time Lapse.” 3. Observe the time course by dragging the slider in the bottom of the “Image Window.” TMRM should fade into noise, while FLIPR brightens up toward the end of the experiment, without out-of-frame dislocation of cells in the image (Fig. 3a, b; see Note 36 and Troubleshooting 7–9). 4. Optionally, match the well labels with condition names. This greatly simplifies pooling technical and experimental replicates when data is recorded to a GraphPad Prism file (see Note 50). This requires that well labels in the image metadata match those of entered into the “dilutions template.xlsx” (see Task #2.3): (a) Click “Edit All Position Names” (list glyph button). (b) Press the “Copy” button. (c) Paste the list into the “dilutions template.xlsx,” “Condition Names” worksheet, from cell A27. (d) Adjust the length of the list in column B if needed. (e) Copy/Paste the list of matched condition names from B27 to the “Multi-Dimensional Open” dialog’s position name list. (f) Save the list. Lists saved using the offered file name and path will be automatically loaded when the recording is opened at any later time.

Millivolts Mitochondrial Membrane Potential in Intact Cells

29

Fig. 3 Typical fluorescence micrographs of FLIPR and TMRM. (a, b) Raw images recorded in H9C2 cells. The images in each channel show baseline, after addition of MDC and after addition of PMPFA. Note the graininess of the images that results from the low light level imaging. Intensity scale bars show gray scale intensity units of a 16-bit cooled, back-illuminated CCD camera. 256
256 pixels, binned images were captured using a 20
lens and are shown without cropping. (c) ROIs overlaid on the processed TMRM channel, first frame. (d) Correct background subtraction is indicated by pixel values fluctuating around zero in cell-free areas of the image (shown light blue in the heatmap scale). (e) Quality control of spectral unmixing. TMRM image corresponding to the end of the recording that was processed with correct, too small and too large FLIPR to TMRM crossbleed coefficient. Arrows indicate (a) gaps in sub-confluent monolayer that are sufficient for background subtraction (b) an area of bright FLIPR fluorescence that was (c) insufficiently removed or (d) left a shadow after spectral unmixing. The black edge marked by (e) is a result of image registration

5. Select the “Mitochondrial membrane potential assay (TMRM/ FLIPR)” or related image analysis pipeline in the main menu “Pipelines/Intensity Measurements/Applications - Membrane Potential.” See Note 37 for choosing the appropriate pipeline for the recording. 6. In the pipeline’s parameter bar (a list at the top left of the main window) adjust values to match the recording (see Note 38). Critical: ensure that the channel number associations are

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correct, and the “ROIs: Cell diameter (pixels)” pipeline parameter matches the average diameter of cells in pixels. 7. Press the “Run Pipeline” (blue double play glyph) button on the main toolbar or in the bottom of the Multi-Dimensional Open dialog. 8. Observe the resultant images (see Fig. 3c–e and Note 39). Critical: if regions of interests (ROIs) are not as desired, adjust the pipeline (see Note 38 and Troubleshooting 3–5) and re-run. 9. At the end of the pipeline execution, the “Membrane Potential Calibration Wizard” appears, with the TMRM and FLIPR fluorescence intensity time course graphs shown in the left panels. 4.10 Task #10: Analysis of Fluorescence Time Courses

The instructions below refer to the use of the “Membrane Potential Calibration Wizard” dialog in Image Analyst MKII. 1. Observe the fluorescence time course traces in the left panels for quality control (QC) and troubleshooting (see Fig. 4a, b, Note 40 and Troubleshooting 10–14). 2. “Calibration Method” tab (see Table 3): (a) Choose the plasma membrane potential calibration method: “Complete with known kP (K-steps) – Goldman using Neural Network.” See alternatives in Notes 41, 42, 46. (b) Choose the mitochondrial membrane potential calibration method: “Complete.” See alternatives in Notes 43– 46. 3. “Wizard” tab; “Data Ranges” sub-tab: (a) Select baseline by holding the left mouse button to select the range in one of the graphs on the left, and press the “Select baseline” button (Fig. 4a, baseline). (b) Select the range after addition of MDC in the graph (the downstroke of TMRM fluorescence and following gradual decay in intensity; Fig. 4b, MDC), and press the “Select MDC (K-eq)” button. Critical: if the downstroke of TMRM is lagging, select from the start of downstroke, or use the “MDC Delay (s)” parameter (see Troubleshooting 22). (c) Select final, complete depolarization in the graph (the plateau after PMPFA addition), and press the “Select CDC (zero)” button (Fig. 4a, CDC). The label “complete depolarization cocktail” (CDC) is used synonymously to PMPFA here.

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Fig. 4 Typical fluorescence time courses of FLIPR (left) and TMRM (right), recorded together using the complete calibration paradigm. (a, b) Correct recording in H9C2 cells, including a single compound addition (the ATP-synthase inhibitor oligomycin) and all calibration steps in Task #8, corresponding to Fig. 3a, b. Gray areas mark frame ranges as selected in the Membrane Potential Calibration Wizard. Arrows indicate key features of the recording: (a) a stable baseline; (b) FLIPR fluorescence is steady during MDC; (c) step-wise increase in FLIPR fluorescence during K+-steps (addition or media replacement with PMK); (d) FLIPR

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(d) Select the K+-steps. To this end, using the FLIPR fluorescence intensity graph, select the end of the MDC segment of the recording, and each segment after PMK applications. Do not include ranges after PMPFA addition. Multiple ranges can be selected. Press the “Select K+ step(s)” button (Fig. 4a, K+-steps). (e) To calculate [K+]ec during K+-steps, enter the K+ concentration in the PM (3.9 mM as given here) and in the PMK (PM + 120 mM ¼ 123.9 mM), and the assay volume before the start of the K+-steps (starting volume + MDC ¼150 μL). Fill the “PMK Removal (ul)” and “PMK Addition (ul)” columns reflecting how the PMK additions were performed in Task #8.15–19. The first line is for the starting [K+]ec corresponding to the MDC, and the corresponding addition and removal fields should be left empty. Press the “Calculate [K+]ec” button. 4. “Wizard” tab; “Constants” sub-tab: set the cell specific parameters here: (a) VF (mitochondria:cell volume fraction): determined in Task #13. Common values range 0.05–0.08 in cell lines. (b) VFM (matrix:cell volume fraction): effects of this variable are largely canceled when using measured aR’ [12]. The default value of 0.8 is used. (c) aR’ (apparent activity coefficient ratio): determined in Task #14. Common values range 0.2–0.5 in cell lines.

ä Fig. 4 (continued) fluorescence reaches plateau after PMPFA addition; (e) well-resolved decay of TMRM fluorescence after MDC addition that starts at lower intensities than the end of the previous segment of the recording; (f) stable, close to zero fluorescence of TMRM at the end of the recording. The following panels help troubleshooting. (c, d) Example recording in immobilized freshly isolated cardiomyocytes from a mouse. Arrows indicate possible problems (a) instability of FLIPR fluorescence during MDC (see Troubleshooting 12); (b) a small overshoot of TMRM fluorescence after MDC addition indicating part quench mode (see Troubleshooting 11). Such recordings need to be repeated at lower TMRM concentration. The unstable TMRM baseline (c) was due to the slow equilibration of TMRM in the large cardiomyocytes, and this is compatible with the calibration as it does not assume equilibrium. (e, f) Example recording in HepG2 cells. Arrows show (a) a small FLIPR response for PMK additions during K+-steps (compare to panels a, c, g) indicating that the specimen is not amenable to complete Nernstian ΔψP calibration using K+-steps (see Troubleshooting 12). A workaround is using the “Complete Iterative” based on the indicated “Part. Depol.” and “CDC” ranges. (g, h) Example recording in human BJ1 quiescent fibroblasts. Minor caveats here are a) too short baseline and (b) non-plateauing CDC segments, that may decrease accuracy. MDC triggered a fast decay in TMRM fluorescence (c) and to capture this, high temporal resolution recording was used (3 s/frame, compared to 30–90 s/frame above) in a single position in order to perform the “Complete” ΔψM calibration in fibroblasts. Colored traces indicate single cells or ROIs. Black traces are mean  SE of all ROIs in a single view field. Fluorescence units (F.U.) correspond to a background-subtracted intensity measured by a 16-bit grayscale camera

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5. Press the “Calibrate” (red play glyph) button to perform the calibration. Typically not all ROIs can be calibrated (see Note 47 on data validation), therefore error messages will appear to note this, unless the “Suppress messages” checkbox in the bottom of the “Wizard” tab is checked. See Note 41 for additional adjustments. 6. Observe the time courses of the calibrated potentials in the right panels for QC (see Fig. 5a, b, Note 48 and Troubleshooting 15–23). 7. Adjust automatic QC parameters (see Note 49) and re-run calibration if needed. This is partly based on estimated error of the calibration and is typically required to reduce noise in addition to the default validation (Note 47). Critical: If calculating single-cell ΔψM, enable QC based on propagated error of baseline (see Fig. 5e and Note 49). 8. Optionally, remove ΔψP traces that did not pass QC for ΔψM, by depressing the “Show only. . .” (PM] glyph) button. 9. Optionally, calculate the mean of traces by setting “Output means” in the “Input/Output” tab or use the context menu of the graph. This is an important data reduction step. The mean is calculated from traces that pass automatic QC, if enabled. 10. To save data, calibrated potentials and fluorescence intensities can be transferred to other software for further analysis by the following ways: (a) Copy/paste graph image or tabular data using the context menu of the graphs. Control the appearance of the graph in the “Input/Output” tab. (b) Configure saving results into Microsoft Excel or GraphPad Prism in the “Input/Output” tab. (c) Fluorescence intensity and calibrated potential traces may be saved in Excel by pressing the button with the Excel icon, and saving the resulting “Excel Data Window” in the main menu “File/Save Excel Data As.” (d) Fluorescence intensity and calibrated potential traces may be added to a GraphPad Prism file, collecting data from multiple conditions and experiments. To this end open or create new Prism file from the main “File” menu, and enable Prism saving in the Membrane Potential Calibration Wizard by pressing the button with the Prism icon (see Note 50). See Note 51 on synchronization and resampling of multiple time courses. 4.11 Task #11 Automated Analysis

1. After setting up the Membrane Potential Calibration Wizard in Task #10, use the save button of the Wizard to save the configuration.

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Fig. 5 Typical calibrated ΔψP and ΔψM time courses. (a, b) Correct recording in H9C2 cells, including a single compound addition (oligomycin) and all calibration steps in Task #8. Fluorescence data corresponding to Figs. 3a, b and 4a, b were calibrated with “Complete with known kP (K-steps) - Goldman using Nonlinear Fit” ΔψP and “Complete” ΔψM calibration algorithms and VF ¼ 0.058 and aR’ ¼ 0.38 values. Data points are mean  SE of 35 calibrated ROIs in the view field, with no additional QC, show as displayed in the Membrane Potential Calibration Wizard. Arrows indicate key features of the recording: (a) a stable baseline; (b) calculated ΔψP not exceeding approximate K+-equilibrium potential (89 mV for 150 mM intracellular [K+] in the used DMEM-formulated PM); (c) potentials for K+-steps are not calculated, because each of these recordings is shorter than the kernel width of the calibration; (d) ΔψM is close to zero and stable after MDC, and (e) remains close to zero after PMPFA. (c, d) Fluorescence traces of HepG2 cells from Fig. 4e, f were calibrated using the “Complete with known kP (K-steps)” (black) or the “Complete Iterative” (red) ΔψP and “Complete” ΔψM calibration algorithms, and VF ¼ 0.078 and aR’ ¼ 0.17 values. Data points show mean  SE of all validated ROIs in the view field with no additional QC. Note that the K+-steps-based complete ΔψP calibration estimated more negative potential, amplified noise (a) and failed in many cells as compared to the iterative method. The ΔψM calibration is the least accurate at depolarized ΔψP and a deviation from zero during

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2. In the main window, “Calibration configuration file name (*. ips)” pipeline parameter, enter the path and filename for the saved membrane potential calibration configuration. 3. In order to automatically save all results either a Prism file must be open (see Task #10.10) or an automatic file naming expression (see main menu “Help/Help on Expression Evaluation”) must be given in the “Output Excel Data save file name (*.xlsx)” pipeline parameter. 4. Using the pull-down menu of the “Run Pipeline” (blue double play glyph) button on the main toolbar or in the bottom of the “Multi-Dimensional Open” dialog run the analysis on all stage positions. 5. Optionally, the “Tools/Batch Pipeline Processor” allows fully automated analysis of many experiments, using one or more calibration configurations and output files. 4.12 Accessory Task #12: Spectral Crossbleed

Spectral crossbleed is determined by calculation from a pair of two-channel images each recorded with only one of the probes present. To maximize fluorescence, TMRM is recorded in baseline condition, whereas FLIPR is recorded with PMPFA to depolarize ΔψP. Prerequisites: an assay plate with cells grown or immobilized in 2 wells, as described for the ΔψM assay (see Note 52). 1. In a 15 mL conical tube, supplement 6 mL 2
PM following the proportions in Table 2 section #1, but omit the probes. 2. In two separate tubes, transfer 2 mL supplemented 2
PM and make PM by adding 2 mL 2
NaCl. 3. In a third tube, take 1 mL supplemented 2
PM and make PMPFA following the proportions in Table 2 section #3. 4. Supplement the two PM tubes either with TMRM or FLIPR using the same concentrations (not critical) as for the ΔψM determination. 5. Supplement the PMPFA tube with FLIPR. 6. Wash a single well with the TMRM-supplemented PM and another well with the FLIPR-supplemented PM, similarly to as given in Task #4, and incubate for minimum 1 h at 37  C ambient air incubator.

ä Fig. 5 (continued) PMPFA addition (b) does not indicate a problem. (e) Fluorescence traces of BJ1 fibroblasts corresponding to Fig. 4e, f were calibrated using the “Complete with known kP (K-steps)” (black) and its “ – Goldman using Nonlinear Fit” variety (red) ΔψP and “Complete” ΔψM calibration algorithms, and VF ¼ 0.042 and aR’ ¼ 0.41 values (aR’ was not measured but from [12]). Data show potentials  estimated SE for each cell in a view field. Using QC based on predicted error of baseline potentials ( second.” This is for TMRM-stained samples. 12. Load images and run the pipeline in each TMRM-stained position. 13. For the parameter “Direction of crossbleed” set “second> first.” This is for FLIPR-stained samples. 14. Load images and run the pipeline in each FLIPR-stained position. 15. In the Excel Data Window average the resultant coefficients. Enter the coefficient matrix into the ΔψM analysis pipeline as “¼{{1,top right coefficient},{bottom left coefficient, 1}}.” 4.13 Accessory Task #13: Mitochondria:Cell Volume Fraction

The VF assay determines the fractional area of mitochondrial to cellular profiles using fluorescence labels in live cells using a laser scanning confocal microscope. If available, electron microscopy stereology values can be also used, expressing the fraction of cellular volume bounded by the mitochondrial outer membrane. Here, use the same specimen as for the ΔψM assay. The use of glass-bottomed vessel is necessary because of the use of oil immersion. 1. Set up an inverted laser scanning confocal microscope with 37  C environment chamber and high NA oil immersion lens to image calcein and MitoTracker Red CMXRos as a two-channel recording 1024
1024 pixels frames at 44 nm pixel size. Use 1 Airy unit pinhole and maximal bit-depth (e.g. 16). Critical: see Task #8.1. Critical: do not use frame averaging or z-stacks. 2. In a 15 mL conical tube, mix 5 mL 2
PM and 5 mL 2
NaCl and supplement following the proportions in Table 2 section #1 for making 1
PM. Include TPB, but omit the probes. Warm to 37  C. 3. In a 2 mL tube pipette 0.3 μL 1 mM calcein-AM and 1.2 μL 100 μM Mitotracker Red CMXRos. The droplets may mix. Rinse 2 mL medium from step #2 on the droplet to disperse. See Note 53.

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4. Wash cell culture in a 1–2 well(s) of cover glass-bottomed vessel 3 times and incubate for 30 min in a 37  C ambient air incubator. 5. Replace fresh, warm medium with probes from step #3 on the cells and transfer the sample to the microscope (see Note 29). 6. In a well area not used for the recording below adjust laser intensities and scan speed. Optimize image for highest quality at slow scanning (see Note 53). At an ideal setting the laser will substantially bleach and damage the scanned view field. Critical: do not saturate detection. 7. Record at least 30 images at different locations, and systematically sampling across the z-dimension, by moving the stage and changing the focus between images. Include planes shearing the bottom or the top of the cells with in focus mitochondria. Critical: Locations do not overlap, because a single scan damages the underlaying cells. Critical: Do not use scanning or epifluorescence with eyepiece to find cells or for focusing. Use bright field instead. Critical: You may center the view field on cells that entirely fit the view field. However, for cells larger than the view filed, it is important to not to bias the recording by centering on a feature e.g. on the nucleus. In this case move randomly in x,y (see Note 54). 8. Open all recorded images corresponding to one sample in Image Analyst MKII using multifile selection. Critical: If the image file(s) open as a “Multi-Dimensional Open” dialog, ensure that all images are loaded as an image series when pressing “Open,” by selecting “Merge in Time” or “Positions as Time Lapse.” 9. Select the “Mitochondria:cell volume fractionator (basic)” image analysis pipeline in the main menu “Pipelines/Morphological Measurements/Applications - Volume Fractionator.” 10. In the pipeline’s parameters set the channel number associations (for other parameters see Troubleshooting 24–25). 11. Press the “Run Pipeline” (blue double play glyph) button on the main toolbar or in the bottom of the Multi-Dimensional Open dialog. 12. Observe the resultant image windows “Cells” and “Mitochondria.” For QC, scroll the image series from the beginning to the end (see Note 55 and Troubleshooting 24–26). 13. Optionally, if dead cells, debris or out-of-focus cells are visible (see Note 55) in the images remove them as follows: (a) Press the Synchronize button on the main toolbar to synchronize frame-to-frame scrolling of the two image channels.

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(b) On each frame, if necessary, draw an ROI around unwanted details, and run the “Mitochondria to cell volume fractionator - mask debris” pipeline. (c) Finally, run the “Mitochondria to cell volume fractionator - recalculate volume fraction” pipeline. 14. Save the resulting “Excel Data Window” in the main menu “File/Save Excel Data As.” The volume fraction is shown in cell G2 as %. Use this value as a fraction in the “Membrane Potential Calibration Wizard,” “Wizard” tab, “Constants” sub-tab. 4.14 Accessory Task #14: Apparent Activity Coefficient Ratio

To calculate the aR’, a confocal microscopic fluorescence time course is recorded following the leakage of TMRM from cells with completely depolarized ΔψM. Then, the ratio of TMRM intensities over the mitochondria-free areas (nucleus) and mitochondrial profiles is calculated. Cell culturing or immobilization in glass-bottomed vessel is necessary because of the use of oil immersion. 1. Follow Task #13.1–2 to set up a laser scanning confocal microscope with the above parameters, but for TMRM imaging, and prepare the assay medium omitting probes and TPB. 2. Add 10
higher concentration of TMRM to the assay medium than using for the ΔψM assay and warm it to 37  C. 3. Wash the cell culture in a few (sample replicates + one setup) wells of coverglass-bottomed vessel 3 times and incubate for 30 min or longer in a 37  C ambient air incubator. 4. In a tube, supplement 0.5 mL assay medium from step #2 with 1 μL MDC stock. Label as MDC. 5. Transfer the sample and the MDC tube to the microscope (see Note 29). 6. Focus cells in the “setup well”, and then half-replace the assay medium with MDC and quickly adjust laser intensities and scan speed. Use a laser intensity and scanning speed that does not result in photo bleaching in multiple scans, but results in a sharp, only slightly grainy image. Critical: do not saturate detection. 7. In a sample well, set laser intensity to the lowest possible level resulting in a faint image of mitochondria in live view and focus into a plane that both intersects nuclei and mitochondria-rich areas. Enable active auto focus if available. If multi-position recording is available, set up 4–8 positions in the same well. 8. Set laser intensity to the level optimized in step #6.

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9. Half-replace the assay medium with MDC in the imaged well and record a time lapse of 10 frames in ~1 min interval (see Note 56). 10. Repeat steps #7–9 in the replicate wells. 11. Open the recording corresponding to one position at a time in Image Analyst MKII. Critical: If the image file(s) open as a “Multi-Dimensional Open” dialog, ensure that when pressing “Open” a time series from single position is opened, e.g. by unchecking “Positions as Time Lapse”. 12. Activate and run the “Mitochondrial membrane potential assay - measurement of the apparent activity coefficient ratio - 1 process pipeline.” 13. In the “Draw nucleus ROIs here” image, draw ROIs over mitochondrion free, nuclear areas (see Note 56). 14. In the “Draw mitochondrial ROIs here” image, draw ROIs over mitochondrion rich areas, corresponding to ROIs drawn in step #13. 15. Activate and run the “Mitochondrial membrane potential assay - measurement of the apparent activity coefficient ratio - 2 – plot” pipeline. 16. In the resultant excel worksheet adjust the range of slope calculation in cell B2 if the graph shows nonlinearity at either ends of the regression analysis. Use the linear part of the curve for regression. 17. Save the data by File/Save Excel Data. The aR’ is shown in cell B2. Use this value in the “Membrane Potential Calibration Wizard,” “Wizard” tab, “Constants” sub-tab.

5

Notes 1. The fluorescence intensity of TMRM is proportional to its concentration under a threshold. When TMRM is used at a concentration below this threshold, the assay is in non-quench mode [6, 23]. In quench mode, much (e.g. 10
) higher concentrations are used and the fluorescence of TMRM is saturated, i.e. “quenched” within mitochondria, where the highest concentrations are reached within the sample. In non-quench mode, a sudden ΔψM depolarization results in decreasing fluorescence intensity. In quench mode this results in a transient spike, followed by a decay of fluorescence. Importantly, for this protocol, TMRM must be used in non-quench mode. 2. FLIPR Membrane Potential Assay Kit contains a proprietary bis-oxonol (FLIPR or PMPI) and an extracellular fluorescence

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quencher. The bis-oxonol is a lipophilic anion and its distribution across the plasma membrane follows chemiosmotic principles. The negative inside ΔψP keeps FLIPR in the extracellular space where its fluorescence is quenched. Therefore, a ΔψP depolarization allows FLIPR to enter the cell and results in an increase of fluorescence. 3. The probes TMRM and the FLIPR Membrane Potential Kit cannot be substituted with other probes, unless kinetic parameters of the plasma membrane redistribution of the new probes are determined. Values we determined for TMRM and FLIPR (see Table 1 in [12]) are pre-loaded in the “Membrane Potential Calibration Wizard,” “Wizard” tab, and “Constants” sub-tab. The FLIPR Membrane Potential Kit is available in blue and red versions. As far as we know, these differ only in the included extracellular quencher, therefore, the biophysical properties of the probe are considered to be the same. We described the technique using the blue version [12], but in our latest publications [18, 19] we used the red version because we observed lower long-term toxicity (unpublished results). The FLIPR dilution is given for the Explorer kit, the bulk version results in 10
more concentrated stocks. 4. The method relies on fluorescence imaging, in order to provide a precise subtraction of background fluorescence intensity that is unsteady in time, and complicated by x,y-positional effects. Therefore, the use of bulk-fluorescence plate readers instead of microscopic imaging is not practical, although is possible [19]. Flow cytometry is another commonly used data acquisition modality to estimate ΔψM. Individual events associated with single cells in a flow cytometer are biased by cell geometry, mitochondrial density, and ΔψP, similarly as with other modalities of fluorescence detection. Without a calibration paradigm performed on the same specimen, it is impossible to completely cancel these confounders. However, cell size and ΔψP can be attributed for by forward scatter and inclusion of a ΔψP probe, respectively, allowing for a better comparison of samples [22]. Importantly, for any flow cytometric ΔψM determination, one needs to be confident that the probe is used in non-quench mode, e.g. by testing it with the application of an uncoupler. 5. The resolution of acquired images should be sufficient to distinguish individual cells. Using a 10
lens is optimal for measuring many cells at a time. A 20
lens is optimal for signal to noise ratio and minimizing photo toxicity. See typical images in Fig. 3a, b. Higher magnifications, where mitochondria are resolved are unnecessary, and will likely cause photo toxicity. Use binning (or configure confocal scanning) to acquire 512
512 pixels images to measure single cells. Use

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256
256 pixels if calculating potentials in clusters of cells. Larger images result in unnecessary image storage and analysis burden, and risk of photo toxicity. 6. To avoid photo toxicity, low light level fluorescence imaging is performed using software-controlled and attenuated light source illuminating the sample only during exposure. 7. Real-time auto-focusing is a critically required component of the instrumentation, in order to obtain a constant focus on the 2–4 h time course in a heated environment with thermal fluctuations caused by liquid handling. In our hands, the Nikon Perfect Focus System on an Eclipse-Ti microscope allowed problem-free recording. A Zeiss Definite Focus on an LSM780 laser scanning confocal microscope (controlled by Zen Black) resulted stable focus only if the entire time course was recorded (with pauses) into a single session. 8. Described media are compatible with ambient air and not with the 5% CO2 environment of a tissue culture incubator. Therefore, any PM-containing assay plate should be incubated in 37  C ambient air incubators, including the heated environment chamber of the microscope. Humidification is not required as long as all reservoir and assay plates are lidded (except see Note 9). 9. Two-photon microscopy requires the use of the red version of the FLIPR Membrane Potential Kit, because the blue quencher absorbs the excitation light leading to excessive heat buildup. If the microscope has upright configuration, the use of microplates and multiple wells described in this method are not possible. Instead, perform the experiment in a single larger vessel (e.g. 35 mm round cell culture dish) and scale up volumes. Mitigating evaporation from this large surface is critical, e.g. by blowing and trapping humidified warm air above the surface. Z-stacking and mean intensity projection may be used for a stable, in-focus recording. 10. Most stocks of fine chemicals, including the MDC stock are stored at 20  C, and repeated freeze-thaw cycles are OK for all listed chemicals. Avoid direct sunlight and over bench or hood illumination, but no other light protection is required if the freezer has an opaque door. 11. The 2
PM may be based on any assay buffer composition, with NaCl and Ca2+ omitted and other component concentrations doubled. Presence of NaHCO3 is optional, and its concentration should not exceed final 5 mM. 12. Filtering prepared media and large volume stocks is recommended to avoid debris interfering with fluorescence imaging. Do not filter media already containing fluorescence probes.

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Another preventable source of debris are pipette tips. Use pre-racked tips and keep racks closed when not in use. 13. Calibration cocktail component stocks may be made by dissolving entire vials of minimal amount of chemicals. The concentrations of these stocks do not need to be precise. 14. Glass vials and inserts are critical for handling PM, because the small, nanomolar concentration TMRM adsorbs on the walls of polypropylene tubes. Of note, we encountered plain glass vials with uncoupler and processed trace clean vials with prooxidant contaminations. The small, 8 mL glass vials are used only for calibration in the presence of mitochondrial inhibitors and uncouplers, so chemical purity is not required. 15. Coverglass-bottomed microplates are recommended (except see Note 9). The potentiometric recording can be also performed in thin plastic bottom microplates at a degraded signal-to-noise ratio and this may introduce problems with auto-focusing. Single-piece polystyrene microplates are not suitable due to their thick bottom. Microplates are recommended in contrast to other microscopy vessels, to allow liquid handling with multichannel pipetting, which is critical when recording fast changes in fluorescence intensity in multiple wells, such as ones evoked by the MDC. 16. Cell cultures below 90% confluency are required, in order to correctly calculate background fluorescence in the cell free areas. Small gaps, such as depicted in Fig. 3a by arrows (a) are sufficient for this. Low cell densities (grown or immobilized) can be used with the assay. 17. Cell cultures adhere less to glass than to the polystyrene material of standard culture vessels. Therefore, we recommend to coat coverglass-bottomed microplates using Matrigel before cell plating. Alternatively, cell type specific coatings may be used. 18. For optimal signal, TMRM concentration should be as high as possible below the threshold of quenching (quench limit, see Note 1). The quench limit can be fundamentally different between different cell types. Because TMRM concentration in the mitochondrial matrix depends on the sum of ΔψP and ΔψM, in cells with very negative ΔψP + ΔψM, e.g. neurons, secretory cells, a low 5–7.5 nM TMRM concentration is recommended. In contrast in hepatocytes (low ΔψP) higher 20–30 nM TMRM is required for a sufficient signal. For each new cell type, determine that application of MDC does not result in a spike of TMRM fluorescence intensity (such as in Fig. 4d), thereby confirming that TMRM is in non-quench mode. The recommended FLIPR concentration is 1:100 or 1: 200 dilution of the reconstituted explorer size vial. Attempt to

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balance TMRM and FLIPR fluorescence so the TMRM intensity at the baseline is similar to the FLIPR fluorescence at the end of the experiment (Fig. 4a, b). To this end, not only concentrations, but illumination intensities and exposure times may be adjusted, and may be different for the two probes. 19. The illumination intensity and exposure time (confocal scan speed) must be optimized in pilot runs. More intense or longer illumination will result in better signal, but with potential photo toxicity. The rule of thumb, here, is to use as much illumination as sufficient for the analysis. This can be accomplished only iteratively, performing the whole analysis and then readjusting. Furthermore, probe concentrations may be adjusted for this (see Note 18). A good starting point is recording visually unpleasing, grainy images (see Fig. 3a, b). Work below the bottom third of the dynamic range of the detector. E.g. in Fig. 3a, b, grayscale fluorescence units were measured up to 1800 on a scale of 0–65,535 (16-bit). Use image intensity histogram in the acquisition software to ensure that the detector is not saturated. Perform a baseline recording, and observe stable intensities and lack of fluctuations (the so-called flickering) in TMRM images. 20. The potentiometric calibration relies on image and data analysis in the software application Image Analyst MKII. It provides a broad range of image file format compatibility, and it is recommended to test first, if the native image format of the microscope system used for the recording can be properly opened. Use a raw, uncompressed or lossless compressed image format, rather than color composite images. Requirements for time stamps are described in Troubleshooting 1. 21. The recommended liquid handling paradigm consists of halfreplacement of the PM each time when test compounds are added after baseline recording. To prevent wash out of previously added drugs, the first addition is prepared at 2
concentrated, the second at 4
, and the third at 8
. Next, the additions are mixed in the prescribed proportions to result in 2
of the final concentration of newly added compounds and 1
of that of compounds already present (see Fig. 2b). To achieve this, mixing volumes are calculated in the “dilutions template.xlsx.” These volumes, shown in rows 83–84 will automatically recalculate if the replacement paradigm is changed in rows 27–28. 22. Preparation of fresh PM and calibrants is recommended on the day of the experiment because of the risk of precipitation, microbial growth (do not filter PM after probes added) and degradation of MDC components during storage. However,

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cleanly prepared, supplemented 2
PM omitting CaCl2 may be stored in a glass container at 4  C for 1 week. 23. The 2
PM stock is prepared with no CaCl2, to prevent precipitation during storage. Calcium phosphate may form a cloudy precipitate in the 2
PM during supplementation if higher concentrations than indicated are used. This is avoidable by supplementing only the final diluted media with CaCl2 or by doing this just shortly before final dilution. 24. Stocks for compound additions are recommended to be 1000–4000
in DMSO. Add DMSO stock first into a dry tube or insert, and then rinse PM on the droplet to disperse. Pipette aqueous or EtOH stocks directly into PM that is already pipetted in the tube or insert. The assay itself is not sensitive to DMSO or EtOH. When using aqueous stocks that are diluted less than 100
, it is recommended to mix an aliquot of the stock solution 1:1 with supplemented 2
PM, and then use this as a half-concentrated stock. In this way potentiometric probes are not diluted by the addition. Alternatively dissolve solids in PM, being mindful of pH changes. 25. The FLIPR component of the assay is incompatible with traces of albumin or serum, therefore these supplements must be avoided. 26. The PMK calibrant is used for stepwise depolarization of the ΔψP. It differs from PM by that 120 mM NaCl was replaced for KCl and is supplemented by MDC stock at final 1
concentration. 27. The PMPFA calibrant serves to completely depolarize ΔψP and ΔψM by application of the Na-ionophore gramicidin and formaldehyde-fixation of the cells. Osmotic support of the cells is critical at this step by the added KCl (Table 2). See “complete depolarization cocktail, (CDC)” in [12] for an alternative formulation that is not formaldehyde-based. 28. The MDC serves as a calibrant that completely depolarizes ΔψM while maintaining ΔψP at a hyperpolarized, K+-equilibrium potential. The main component is the K+-ionophore valinomycin that serves both of these purposes. Other core components help depolarization of ΔψM. In addition, MDC has optional components that mitigate cell swelling (see Table 1, Note 47), and their use may depend on the cell type. Generally, these optional components are required if the complete ΔψP calibration reports larger MDC-triggered hyperpolarization in their presence, than when using MDC only with its core components. MDC contains MDC stock at 3
of final concentration. 29. Avoid cooling down the culture plate while liquid handling or while transferring between facilities. Avoid placing culture

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plates on cold surfaces, like benchtops, TC microscopes before the assay. Carry plates sandwiched between warm bricks. 30. The incubation of the sample in PM, followed by the “final wash” (Task #7) serves to equilibrate both the specimen and the sidewalls of the microplate with TMRM, and to wash out traces of albumin that interferes with the assay. The PM for the “final wash” is handled identically to the additions in reservoirs with glass inserts, so that subsequent media replacements do not change probe concentrations. 31. The cycle time of the instrument, thus, how long it takes to record all desired positions or wells, is a critical determinant of the experimental design. To perform a successful “complete calibration,” the decay in TMRM fluorescence after complete depolarization of the ΔψM needs to be resolved by at least 15 frames. In large cells, this may work with 30–90 s cycle time and 30 min recording after MDC addition (Fig. 4b, d, f). Cells with large surface to volume ratio will show faster changes in TMRM fluorescence intensities and need shorter cycle time (e.g. 3 s in Fig. 4h). Longer cycle times, and therefore extension of recordings to larger numbers of wells, can be used with alternative calibrations (see Notes 42–43). 32. There is no requirement for an electrochemical equilibrium of the potentiometric probes at start of the baseline, because calibration equations account for disequilibrium. However, to avoid an x,y- or focal drift, the assay plate and the heated microscope enclosure (environment chamber) needs to be properly thermally equilibrated that warrants for proper equilibration time on the microscope stage. Furthermore, to help image registration, before each segment of recording, or after pipetting, push the microplate by fingers into the opposite corner from the retaining spring. 33. The minimum number of frames in a segment of recording equals the differentiation kernel width in the Membrane Potential Calibration Wizard “Constants” tab, which is 7 at default. This will result in a single time point of calibrated potential. Each additional frame will result in an additional calibrated data point. A longer baseline recording improves the accuracy of determination of basal potentials. 30 min or 30 frames, whichever is shorter in time is recommended for baseline recording. 34. Record each segment of the experiment (e.g. baseline, after addition #1. . .) into separate recordings, thus stopping rather than pausing acquisition after each segment, if practical. This works well on Nikon Elements software, but in our hands caused problems by resetting auto-focusing on a Zeiss LSM

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system (Zen Black), and this latter case pausing the time course for additions worked well. 35. The last segment of the experiment has to be sufficiently long to capture a plateau in the FLIPR fluorescence intensity. This is common for all calibration paradigms. In some cell types it may be extended longer than the recommended 30 min. After the recording is complete, the sample has been fixed in PFA and is amenable to immunocytochemistry. FLIPR and TMRM wash out during immunostaining. In certain microscope systems the x,y-coordinates are saved as image metadata and can be later be used to find the cells from the live cell recording, image immunofluorescence at the same positions, and gate the potentiometric analysis to a specific cell type. To this end, use “with post-hoc ICC” image processing pipeline version for analysis in Image Analyst MKII. 36. Visual inspection of raw recordings is useful for QC and troubleshooting. Images opened in Image Analyst MKII are intensity auto scaled by default. Therefore, loss of intensity appears as brightening background noise (Fig. 3a). Expect bright TMRM fluorescence at the baseline, in a visibly mitochondrial pattern, clustered in the perinuclear region. TMRM fluorescence should show no fluctuations, otherwise this may indicate too high level of illumination. At latest, following MDC addition TMRM fluorescence must fade out (see Troubleshooting 7–9). Typically, the mitochondria to cytosol translocation of TMRM is observable as TMRM images suddenly appear blurry and any discernible mitochondrial pattern is lost. FLIPR fluorescence is typically dim at baseline, but may be bright in cell lines with a low ΔψP, such as hepatocytes. FLIPR fluorescence should stepwise increase during PMK additions, reaching a stable high intensity after PMPFA. Shaking of the view field and occasional floating in debris are typical, and cause no problem for the analysis. 37. Raw fluorescence image recordings are processed by channel alignment (to correct pixel shift by filter sets), image registration (to correct x,y-misalignment by stage motorization), background subtraction (to eliminate electronic offsets and probe fluorescence in the medium), and spectral unmixing in Image Analyst MKII. Finally, the pipeline performs automated ROI drawing, by recognizing single or groups of cells. Fluorescence intensity time courses are further analyzed in the “Membrane Potential Calibration Wizard.” While pipelines are editable, different configurations are provided for major experimental design scenarios. The prototype pipeline “Mitochondrial membrane potential assay (TMRM/FLIPR)” was designed for wide field microscopy and single-cell selection, and it implements all of the above image processing steps. The “. . .

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for CONFOCAL” variants were simplified by removal of channel registration and background subtraction. If cells move considerably during the time course, it is recommended to calculate ΔψM over groups of cells. The “. . . for groups of cells” pipeline variants will outline clusters or colonies of cells. If low baseline cell viability compromises the assay, cells with depolarized ΔψP (indicated by high FLIPR fluorescence) can be automatically avoided by using the “. . . with masking dead cells” pipeline variants. The “. . . with post-hoc” pipeline variants allow using a separate immunofluorescence or bright field recording to gate single-cell analysis to a specific cell population. 38. Pipeline parameters “Channel Number, TMRM” and “Channel Number, FLIPR” are the ordinal numbers of the respective channels in the recording. “Local background subtraction by median rolling ball” toggles rolling median background subtraction using a window width provided in “Rolling ball: Median filter size (pixels).” The window width should be considerably larger than the diameter of the cells. If rolling ball is disabled, the pixels that are darker for the entire time course than a percentile specified in “Background level (percentile)” are used to calculate the background level in each frame and channel. Typical background level settings are: 5 percentile for near confluent, 20 for sub-confluent, and 50 for very sparse cultures. The same pixel positions are used for the entire time course in order to provide a background subtraction that is consistent in time. The “Spectral Unmix Coefficient Matrix” provides the spectral crossbleed matrix in the following form: “¼{{1, fractional crossbleed of FLIPR into TMRM channel},{fractional crossbleed of TMRM into FLIPR channel, 1}}, and is unrelated to the recording order of the channels (see Task #12) for determination of these values. The image stabilizer operates on the sum of the two channels, and the parameters below control how time courses are registered in x,y. “Image stabilizer compares to first frame” toggles if frames are registered by comparing each frame to the first frame, or on a rolling-basis using “Image stabilizer period (frames)” number of frames around each frame to calculate dislocations. The next set of parameters control ROI drawing. Because this assumes cell size, the following parameters likely need to be tuned for the specific sample and microscope configuration. ROIs are generated on a maximum intensity projection of the time course of the sum of the two channels. Therefore, these ROIs will attempt to capture the complete dwelling area of slightly motile cells. We have previously shown that an exact way of ROI drawing, including more background around a cell, does not bias the results [12]. Cells with

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diameter broadly ranging around “ROIs: Cell diameter (pixels)” will be selected. If single cells are detected as multiple ROIs, increase this value. If multiple cells are detected as a single ROI or larger cells are missed, decrease this value. Another toggle controlling splitting objects into multiple ones is the “ROIs: weld segments into round objects.” Cell boundaries will be extended by “ROIs: Margin around cells (pixels)” amount. Increasing this value helps to encircle the area where a motile cell dwells during the recoding. The detection of cells is also controlled by “ROIs: Minimum Cell Fluorescence (%).” Increase this value if background is circled instead of cells, or decrease this value if dimmer cells are missed. To avoid bright debris or hot pixels interfering with ROI drawing, automatic scaling is performed compared to a slightly less than 100 percentile value of the image histogram “ROIs: Debris cutoff (percentile).” This percentile value defines the 100% for the minimum fluorescence parameter above. Decrease this value to make ROI drawing more sensitive and enter a value closer to 100 to make it less sensitive to detect objects. Notably, if bright large cells are missed, increase the cell diameter value above. The “ROIs: Sensitivity of cell boundaries (% of max fluorescence)” parameter controls how a cell is circled, compared to its local, maximal fluorescence. A smaller value will result in larger ROIs. The last four pipeline parameters control data processing. In “Membrane potential calibration action” select an action to be performed by the Membrane Potential Calibration Wizard, by choosing between loading data only, or also performing calibration. The latter is required for automated operation. “Calibration configuration file name (*.ips)” is left blank for the first run of the pipeline. Calibration configuration files can be saved in the Membrane Potential Calibration Wizard. It is recommended to use a dedicated calibration configuration file for each experiment, as this contains the information on the timing of the calibrant additions. Optionally, in the “Output Excel Data save file name (*.xlsx),” provide a file name, or an automatic file naming expression (see main menu Help/Help on Expression Evaluation) if a Microsoft Excel output is preferred. Excel output is per view field. Alternatively, see Note 50 on storing multiplexed data in a GraphPad Prism file. 39. Visual inspection of processed images is important while setting up the assay. Shaking of the view field when playing the time lapse should be absent (instead, edge areas may be missing and shown in black; see Troubleshooting 3). Moving the pointer over dark areas of the image should indicate intensities fluctuating around zero (positive and negative values, Fig. 3d). To verify spectral unmixing, observe whether the

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baseline is free of negative (dark) imprints of TMRM fluorescence in the FLIPR channel, and the end of the recording is devoid of such imprints of FLIPR fluorescence in the TMRM channel (Fig. 3e). 40. Visual inspection of fluorescence intensity traces is important while setting up the assay (Fig. 4a, b). First inspect the baseline stability. While an electrochemical equilibrium of the probes is not required for the calibration, unstable baseline may indicate problems with the sample (Troubleshooting 6–8). FLIPR fluorescence may decrease (ΔψP hyperpolarization) or increase (ΔψP depolarization) during MDC addition. This is dependent on the cell type. Successful complete calibration paradigm requires at least partially maintained ΔψP during the MDC phase of the recording. FLIPR fluorescence must show a plateau at the end of the recording. It may be preceded by a smaller overshoot, but it should not precipitously drop (Troubleshooting 12–14). The TMRM intensity at the end of the recording is expected to be stable and slightly above zero. If this is not the case, see Troubleshooting 4 of spectral unmixing. 41. The complete ΔψP calibration (“Complete with known kP (K-steps) - Goldman using Neural Network”) determines calibration points using the Goldman equation. This paradigm requires establishing K+ as a major or partially ΔψP-governing ion using the K+-ionophore-containing MDC, stepwise increase of [K+]ec and finally complete depolarization of the ΔψP. It calculates four parameters, ΔψP at the baseline, FLIPR fluorescence at zero potential (fP0), cellular background fluorescence (including probe binding and autofluorescence; fPX), and the contribution of ionic conductances other than K+ to the ΔψP during calibration (PN, see Eq. 9 in [12]). The neural network-based calibration may require adjustment of the network training ranges for the above parameters in the “FLIPR Parameters” tab. The neural network is automatically trained when needed by modeling FLIPR fluorescence based on Goldman equation and the given [K+]ec steps. Because typically residual ion conductances contribute to the ΔψP in the presence of MDC, the Goldman equation-based calibration is recommended. However, in specimens where this effect is small (PN < 5) use the more robust, linear fit-based “Complete with known kP (K-steps)” method. This uses Nernst equation when PN ¼ 0. The estimated PN value can be found in “Wizard” tab, “Assumed and Calculated Parameters” sub-tab or in the parameter list section of saved data. kP is a rate constant that is not measured in typical applications and a previously determined value, 0.38 s1 is used [12]. Determination of kP is required only for high temporal resolution (e.g. 1 s frame

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interval) recordings, and may be performed using the “Complete (temporally resolved K-steps)” method. Alternative calibrations below allow for a simplified experimental design by re-using previously measured, or published baseline ΔψP (Note 42) or by using the iterative algorithm (Note 46). 42. The “Baseline & Zero” ΔψP calibration allows for a simplified experimental design by re-using previously measured, or published baseline ΔψP. This requires only establishing zero potential at the end of the assay by the addition of the PMPFA, while no MDC or PMK additions are used. ΔψP is calibrated by a 2-point calibration in this case, to the baseline and the zero value. Variations for this approach are the “Baseline & K-equilibrium” and “K-equilibrium & Zero” calibration paradigms, where “K-equilibrium” refers to a ΔψP calculated assuming K+-equilibrium potential and intracellular [K+] during addition of the K-ionophore containing MDC. Before calibration, provide required parameters in the “Wizard” tab, “Assumed and Calculated Parameters” sub-tab. 43. The complete ΔψM calibration (“Complete”) determines four parameters: ΔψM at the baseline, TMRM fluorescence at zero potential (fT0), and cellular background fluorescence (probe binding and autofluorescence; fTX), and the rate constant of TMRM redistribution across the plasma membrane at zero potential (kT). This paradigm requires a sudden and complete depolarization of ΔψM that results in exponential-like decay in TMRM fluorescence intensity during the course of several minutes, followed by complete depolarization of the ΔψP as well. It is therefore critical, that ΔψM is significantly polarized (about TMRM” checkbox, re-running the analysis, and adding the residual crossbleed coefficient to the “top right coefficient.” The TMRM to FLIPR crossbleed is typically small, and it can be measured only in single-probe stained samples as described in Task #12. 5. ROIs are not outlining cells as desired. Follow instructions in Note 38 to optimize ROI drawing parameters of the pipeline. 6. Baseline fluorescence is not steady. This is not necessarily a problem, but may indicate too short incubation time, high illumination intensity and photo toxicity, incorrect temperature in the environment chamber or the result of cold transportation of the sample between incubator and the environment chamber. 7. TMRM fluorescence is weak, FLIPR fluorescence is strong on baseline. Consider cell viability problems. If the ΔψP is known

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to be small (depolarized) in the sample, TMRM concentration needs to be increased, and FLIPR concentration may be decreased. 8. TMRM fluorescence flickers during baseline recording. Decrease illumination intensity or exposure times and restart recording in a new, previously unilluminated location. 9. TMRM fluorescence does not fade out during MDC addition. Pipette the MDC closer to the cell monolayer and mix during MDC addition more vigorously. Asynchronous response of cells in a wave across the view field typically indicates insufficient mixing. Lipofuscin autofluorescence or microbial contamination may also lead to sustained fluorescence in the TMRM channel in a punctate pattern reminiscent of mitochondria. These don’t interfere with the calibration only as long as these intensities are also observed during the PMPFA recording, because the calibration algorithm will attribute it to auto- or bound probe fluorescence (fTX). In such case, the TMRM’s CDC range may be taken as the beginning of the recording after PMPFA application, ensuring that the contaminating fluorescence is captured without a loss. 10. TMRM fluorescence intensity drops step-like after MDC, and no decay curve can be observed. The temporal resolution of the recording (including cycle time of the multi-position recording, and the speed of restarting recording after pipetting) is insufficient to capture the decay. This may happen for cell types with large surface to volume ratios, small mitochondrial content or low ΔψM. Consider recording fewer positions and wells to shorten pipetting and recording cycle time. Consider using higher cell density (will decrease surface to volume ratio) or higher TMRM concentration (if can be increased in non-quench mode). Alternatively, use “Complete (known k)” calibration method for ΔψM, which does not rely on capturing the decay curve (see Note 44). Alternatively, determine baseline ΔψM in a single well at a higher temporal resolution using the “Complete”, and then use “Baseline & Zero” calibration in larger numbers of wells recording at a lower temporal resolution (see Notes 42–45). 11. TMRM fluorescence intensity overshoots at the application of MDC and this is followed by a decay (Fig. 4d). The overshoot indicates that TMRM was in quench mode (see Note 1) and this results in inaccurate calibration. Repeat the experiment with decreased TMRM concentration. 12. FLIPR fluorescence increases during MDC, and/or it is not or very little responsive to PMK or PMPFA additions (Fig. 4c, e). The specimen may not be amenable for complete calibration (see Note 40). First, try to decrease valinomycin concentration

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and omit FCCP and optional components in the MDC. Alternatively, use the iterative (see Note 46) or the baseline to zero calibration for ΔψP (see Note 42) with an assumed baseline potential. In this case ΔψP is likely to be small, and errors in ΔψM made by assuming ΔψP will be small. 13. FLIPR fluorescence intensity rises and gradually decays after each PMK addition. It is likely to be because of excessive ΔψP depolarization and then recovery during PMK addition. Pipette or mix more gently, and away from the cell layer, so high K+ PMK does not directly hit the cells. Increase the wait time before restarting image acquisition. 14. FLIPR fluorescence rises and then decays during PMPFA application. Verify if PFA was correctly pH-ed and all components of PMPFA were in place. Increasing osmolality of PMPFA may help to avoid the decay. Alternatively use CDC defined in [12]. 15. No potential traces appear after calibration. Verify range selections in the “Wizard tab.” Check the status bar of the Membrane Potential Calibration Wizard for an error message. Turn off “Suppress messages” in the Wizard tab, and re-run calibration. Then, troubleshoot ΔψP calibration by trying alternative methods on the same data (see Notes 41, 42, and 46). For further troubleshooting check mark the Wizard tab, Expert mode, “Edit all parameters.” Relax validation criteria “r^2 Min” values at “FLIPR Parameters” and “TMRM Parameters” tabs. Disable “Apply point-by-point range criteria” in the “Constants tab.” Furthermore, disable QC toggles in “FLIPR Parameters” and “TMRM Parameters” tabs. Note: a fatal error in calibration may originate from wrong frame range settings, or insufficient number of frames for the MDC segment, or from problems at data export. 16. ΔψM cannot be calibrated because of too short MDC frame range. Ensure that the selected MDC frame range is at least 2 frames longer than the kernel width “Differential kernel width” in the “Constants” tab. The useful frame range may be too sort, because the actual range that is used for calculation with the TMRM fluorescence intensity decay is truncated to eliminate initial high rate of fluorescence drop and trailing frames where the temporal derivative fluctuates around zero. Control the initial cut off by increasing the “TMRM Parameters” tab “Maximum allowed DTE” value, or the trailing cut off by the “Range of regression ends at zero DTE.” DTE refers to the normalized temporal derivative of TMRM fluorescence intensity [12], and can be visualized by pressing the “ΔψM” button on the left of the potentials graph. Alternatively, decrease the kernel width or record longer after

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application of MDC. The “Wizard” tab, “Expert mode,” “Edit all parameters” must be check marked to access kernel width. Alternatively use “Complete (known k)” calibration paradigm for ΔψM (see Note 44). 17. ΔψM calibration works in control condition but not after a specific compound addition. A test compound may interfere with the complete calibration paradigm by discharging ΔψM before calibration or by altering plasma membrane properties. In such cases, measure the value of kT in control condition using complete calibration and use the “Complete (known k)” method, or also measure baseline ΔψP and ΔψM, and then re-evaluate all conditions using the “Baseline & Zero,” “Baseline & MDC or CDC” methods. 18. Calibrated potentials are too noisy. If the temporal resolution allows, greater noise suppression can be achieved by increasing the “Differential kernel width” in the “Constants” tab. Increasing exposure intensities or times during recording, or the frame rate of the recording can help to increase signal to noise ratio, but may cause photo toxicity. Using a high NA 20
lens instead of 10
magnification, is a good way of increasing signal-to-noise ratio. Compare fluorescence time courses plotted in raw and processed images to determine if image registration or background subtraction injects noise, and adjust their settings, see Note 38. 19. Few of the ROIs result in calibrated potentials or the estimated error or variability of single-cell baseline potentials is very large. As a rule of thumb, a calibration paradigm fitting fewer parameters on the data will result in smaller estimated errors and a larger fraction of ROIs passing validation and QC. To this end, consider performing a two-step calibration, where first a cell population average of one or more of the following parameters are determined: kT, PN, baseline ΔψP and ΔψM, depending on which of these can be assumed to be uniform in the cell population. Then in a second-pass calibration, use the determined parameter(s) and the respective calibration method (“Complete (known k),” “Complete with known kP (K-steps),” “Baseline & Zero,” or “Baseline & MDC or CDC”) to calculate single cell potentials with a smaller cellto-cell error. 20. Calibrated ΔψP is more negative than expected during baseline or during MDC application. For this type of calibration error, it is diagnostic if the calibrated ΔψM shows substantial polarization after PMPFA. This may happen when using the Nernst equation-based “Complete with known kP (K-steps)” method, and the K+-equilibrium potential is not properly established during K+-steps calibration. The K+-equilibrium potential

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during K+-steps dictates a linear relationship between [K+]EC-zp and FLIPR fluorescence (see details in [12]), and a linear regression for these data is used for calculation of ΔψP calibration parameters. Press the ΔψP button in the Membrane Potential Calibration Wizard to show this regression plot. Data not well fit with a line indicate the non-Nernstian behavior of ΔψP, thus that K+ permeability does not govern ΔψP precisely. In such a case, use the “Goldman” variant of the ΔψP calibration with the standard, complete calibration paradigm involving K+-steps or the “Iterative” paradigm using no PMK additions. The “Goldman” variant performs a non-linear fit on these data assuming significant contribution of other ionic permeabilities, or ion pumping to the generation of ΔψP. Alternatively, increase valinomycin concentration in the MDC and omit FCCP. 21. ΔψM cannot be calibrated, but calibrated ΔψP traces are shown. First, turn off QC (see Troubleshooting 15) to see if QC rejected the traces. Overestimation (too negative) ΔψP during MDC can cause this, see Troubleshooting 20. To troubleshoot “complete” or “complete (known k)” ΔψM calibrations, press the “ΔψM” button on the left of the potentials graph. This shows the DTE vs FTE plots (defined in [12] and see Troubleshooting 16). If no traces appear in this plot, it can be due to no valid calibrated ΔψP points for the MDC range or due to wrong frame range settings for ΔψM calibration. If traces appear, the center part of the DTE vs FTE plots should be linear for each ROI (marked by a different color). A deviation from linear at the high DTE values may indicate too slow action of MDC. Exclude these high value points by a lower “Maximum allowed DTE” value in the “TMRM Parameters” tab or by a larger “MDC Delay (s)” value. Mix more vigorously upon MDC addition or increase valinomycin or FCCP concentrations. This is a parametric plot and a curly or wavy pattern may indicate insufficient mixing after MDC addition or positioning/focusing instability. Alternatively, troubleshoot (4) spectral unmixing. 22. ΔψM is not zero after MDC. Verify the selection of the MDC range. If the decay in TMRM fluorescence starts with a delay after MDC addition, use a “MDC Delay (s)” value of 60–90 s in the “TMRM Parameters” tab. If this affects a sub-population of cells, reject them by QC, using the “Quality control based on MDC calibrates to zero” toggle in the “TMRM Parameters” tab. If zero is not reached in a step-like fashion in the calibrated ΔψM trace during MDC, but ΔψM changes gradually or overshoots to positive values, then it is likely that the value of kT was not determined accurately or the

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MDC triggered only gradual depolarization of ΔψM. See Troubleshooting 21 of DTE vs FTE plots to diagnose this. 23. ΔψM is not zero after PMPFA. The ΔψM calibration is the least accurate when the ΔψP is fully depolarized [12]. A smaller deviation from zero (~50 mV) does not indicate a problem. A consistent deviation from zero may indicate inaccurate ΔψP calibration, and using alternative ΔψP calibration paradigms is recommended (see Troubleshooting 20 and Notes 42, 46). 24. In the VF analysis, most frames disappear. The pipeline parameter “Minimal mean image intensity cutoff for the cytoplasmic stain” is used for automatic removal of frames that contain too few details. Decrease this cutoff value. 25. In the VF analysis, the “Cell” image does not show profiles of whole cells. If calcein accumulates in subcellular structures, and only these structures are visible, decrease the “Sensitivity scaling for the cytosolic stain (percentile)” value the pipeline parameters and re-run analysis in until the profile for the whole cell is shown. If background is included and cell profiles have fuzzy edges, increase this value (up to 100). 26. In the VF analysis, the “Mitochondria” image shows mitochondrial profiles with fuzzy edges. The quality of images needs to be improved, by increasing MitoTracker concentration or slowing scanning.

Acknowledgments The authors thank Kareem Heslop (Hollings Cancer Center, Medical University of South Carolina, SC. USA) for critical reading of the manuscript, Christopher Wiley and Leyla Teos (Buck Institute for Research on Aging, CA, USA) for providing cultures of BJ1 cells and freshly isolated cardiomyocytes, respectively. This work was supported by the National Institutes of Health (NIH) grants 1R41DA043369 to AAG and R01AG055822 to Simon Melov. AAG declares a financial interest in Image Analyst Software. References 1. Brand MD (1995) Measurement of mitochondrial protonmotive force. In: Brown GC, Cooper CE (eds) Bioenergetics: a practical approach. IRL Press, Oxford, pp 39–62 2. Sekine S, Youle RJ (2018) PINK1 import regulation; a fine system to convey mitochondrial stress to the cytosol. BMC Biol 16:1–12. https://doi.org/10.1186/s12915-0170470-7

3. Schendzielorz AB, Schulz C, Lytovchenko O et al (2017) Two distinct membrane potentialdependent steps drive mitochondrial matrix protein translocation. J Cell Biol 216:83–92. https://doi.org/10.1083/jcb.201607066 4. Palmieri F (2013) The mitochondrial transporter family SLC25: identification, properties and physiopathology. Mol Asp Med 34: 465–484. https://doi.org/10.1016/j.mam. 2012.05.005

Millivolts Mitochondrial Membrane Potential in Intact Cells 5. Kim SJ, Xiao J, Wan J et al (2017) Mitochondrially derived peptides as novel regulators of metabolism. J Physiol 595:6613–6621. https://doi.org/10.1113/JP274472 6. Nicholls DG (2012) Fluorescence measurement of mitochondrial membrane potential changes in cultured cells. Methods Mol Biol 810:119–133 7. Zorova LD, Popkov VA, Plotnikov EY et al (2018) Mitochondrial membrane potential. Anal Biochem 552:50–59. https://doi.org/ 10.1016/j.ab.2017.07.009 8. Lemasters JJ, Ramshesh VK (2007) Imaging of mitochondrial polarization and depolarization with cationic fluorophores. Methods Cell Biol 80:283–295 9. Brand MD, Nicholls DG (2011) Assessing mitochondrial dysfunction in cells. Biochem J 435:297–312. https://doi.org/10.1042/ BJ20110162 10. Choi SW, Gerencser AA, Nicholls DG (2009) Bioenergetic analysis of isolated cerebrocortical nerve terminals on a microgram scale: spare respiratory capacity and stochastic mitochondrial failure. J Neurochem 109:1179–1191. https://doi.org/10.1111/j.1471-4159.2009. 06055.x 11. Merlini L, Angelin A, Tiepolo T et al (2008) Cyclosporin a corrects mitochondrial dysfunction and muscle apoptosis in patients with collagen VI myopathies. Proc Natl Acad Sci U S A 105:5225–5229. https://doi.org/10.1073/ pnas.0800962105 12. Gerencser AA, Chinopoulos C, Birket MJ et al (2012) Quantitative measurement of mitochondrial membrane potential in cultured cells: calcium-induced de- and hyperpolarization of neuronal mitochondria. J Physiol 590: 2845–2871. https://doi.org/10.1113/ jphysiol.2012.228387 13. Keij JF, Bell-Prince C, Steinkamp JA (2000) Staining of mitochondrial membranes with 10-nonyl acridine orange, MitoFluor green, and MitoTracker green is affected by mitochondrial membrane potential altering drugs. Cytometry 39:203–210. https://doi.org/10. 1002/(sici)1097-0320(20000301)39: 33.0.co;2-z 14. Kholmukhamedov A, Schwartz JM, Lemasters JJ (2013) Isolated mitochondria infusion mitigates ischemia-reperfusion injury of the liver in rats: mitotracker probes and mitochondrial membrane potential. Shock 39:543. https://

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doi.org/10.1097/01.shk.0000430660. 63077.7f 15. Gerencser AA, Mookerjee SA, Jastroch M, Brand MD (2016) Measurement of the absolute magnitude and time courses of mitochondrial membrane potential in primary and clonal pancreatic beta-cells. PLoS One 11:e0159199. https://doi.org/10.1371/journal.pone. 0159199 16. Nicholls DG (2006) Simultaneous monitoring of Ionophore- and inhibitor-mediated plasma and mitochondrial membrane potential changes in cultured neurons. J Biol Chem 281:14864–14874 17. Gerencser AA (2015) Bioenergetic analysis of single pancreatic β-cells indicates an impaired metabolic signature in type 2 diabetic subjects. Endocrinology 156:3496–3503. https://doi. org/10.1210/en.2015-1552 18. Gerencser AA (2018) Metabolic activationdriven mitochondrial hyperpolarization predicts insulin secretion in human pancreatic beta-cells. Biochim Biophys Acta Bioenerg 1859:817–828. https://doi.org/10.1016/j. bbabio.2018.06.006 19. Gerencser AA, Mookerjee SA, Jastroch M, Brand MD (2017) Positive feedback amplifies the response of mitochondrial membrane potential to glucose concentration in clonal pancreatic beta cells. Biochim Biophys Acta Mol basis Dis 1863:1054–1065. https://doi. org/10.1016/j.bbadis.2016.10.015 20. Birket MJ, Orr AL, Gerencser AA et al (2011) A reduction in ATP demand and mitochondrial activity with neural differentiation of human embryonic stem cells. J Cell Sci 124:348–358. https://doi.org/10.1242/jcs.072272 21. Guntur AR, Gerencser AA, Le PT et al (2018) Osteoblast-like MC3T3-E1 cells prefer glycolysis for ATP production but adipocyte-like 3T3-L1 cells prefer oxidative phosphorylation. J Bone Miner Res 33:1052–1065. https://doi. org/10.1002/jbmr.3390 22. Rovini A, Heslop K, Hunt EG et al (2021) Quantitative analysis of mitochondrial membrane potential heterogeneity in unsynchronized and synchronized cancer cells. FASEB J 35:100–106. https://doi.org/10.1096/fj. 202001693R 23. Nicholls DG, Ward MW (2000) Mitochondrial membrane potential and cell death: mortality and millivolts. Trends Neurosci 23:166–174. https://doi.org/10.1016/S0166-2236(99) 01534-9

Chapter 3 Rotenone Decreases Ischemia-Induced Injury by Inhibiting Mitochondrial Permeability Transition: A Study in Brain Ramune Morkuniene, Evelina Rekuviene, and Dalia M. Kopustinskiene Abstract Mitochondria participate in many physiological and pathological processes in the cells, including cellular energy supply, regulation of calcium homeostasis, apoptosis, and ROS generation. Alterations of mitochondrial functions, especially the opening of mitochondrial permeability transition pore (mPTP) are the main mechanisms responsible for the ischemic brain damage. Recently, the inhibitors of the Complex I of mitochondrial respiratory chain emerged as promising suppressors of mitochondrial ROS generation and mPTP opening. Here we describe the assay that can be implemented easily to evaluate the protective effects of rotenone or other potential inhibitors of the Complex I of mitochondrial respiratory chain against acute ischemia-induced injuries in brain. Key words Rotenone, Brain ischemia, Isolated brain mitochondria, Mitochondrial respiration, Calcium retention capacity, Mitochondrial permeability transition pore, Complex I, Complex II, ROS generation

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Introduction The restoration of normal mitochondrial functions and inhibition of mitochondrial permeability transition pore (mPTP) could be one of the most promising strategies to decrease the brain damage during ischemia [1, 2]. The possibility to use known inhibitor of mPTP cyclosporine A [3, 4] is limited due to its impaired transport into the brain [5], low efficacy in brain mitochondria [6], and neurological side effects [7]. However, recently inhibitors of the Complex I of mitochondrial respiratory chain, e.g. rotenone [8], metformin [9], and amobarbital [10] were suggested as potential agents to inhibit mPTP, suppress ROS generation, and reduce cell death. Here we propose assays routinely used in our investigations to determine whether Complex I of the mitochondrial electron transfer system may be involved in regulation of mPTP opening during ischemia and whether inhibitors of Complex I—rotenone or

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other compounds can protect against ischemia-induced cell death in an experimental model of total ischemia in adult rat brains. The successful lead compound, after the intravenous injection 20 min before ischemia, should increase the resistance to Ca2+-induced mPTP opening, increase the mitochondrial respiration rate, and decrease production of ROS in mitochondria isolated from ischemia-damaged cortex and cerebellum [11–13].

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Materials Important—for all experiments with isolated mitochondria, double-distilled water (the second time—using a glass distillation system) should be used instead of ultrapure water to prepare all solutions and media. Ion-exchange resins used in water purification systems are potent inhibitors of mitochondrial functions therefore ultrapure water should be avoided in these types of experiments. 1. Four male Wistar rats (8–12 weeks, weight 200–250 g). 2. Vehicle: ethanol/0.9% NaCl, 1:5. 3. Isolation medium: 225 mM mannitol, 75 mM sucrose, 5 mM HEPES, 1 mM EGTA, pH 7.4 at 4
C. 4. Hypoxic gas mixture (93% N2, 5% CO2, 2% O2). 5. Incubation buffer: 200 mM sucrose, 10 mM Tris–HCl, 1 mM KH2PO4, 10 μM EGTA, pH 7.4 at 25
C. 6. Respiration medium: 110 mM KCl, 10 mM Tris–HCl, 5 mM KH2PO4, 2.24 mM MgCl2, pH 7.2 at 37
C (for respiration measurements) or at 25
C (for H2O2 generation assessment). 7. 25 mM KH2PO4 buffer (pH 7.4 at 25
C). 8. 50 mM KH2PO4 buffer (pH 7.4 at 25
C).

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3.1 Animals and Experimental Protocols for Rotenone Treatment and Brain Ischemia

1. Randomly assign rats to four experimental groups: control, rotenone control, 120 min ischemia, rotenone plus 120 min ischemia. 2. For the evaluation of rotenone effects, slowly infuse vehicle or rotenone at a dose of 0.1 mg/kg in 0.2 mL of vehicle into the tail vein of rats. 3. After 20 min, expose the rats from all groups to increasing concentrations of CO2 in the air in CO2 euthanasia chamber and then sacrifice the rats by cervical dislocation and decapitation.

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4. Remove the brains using scissors, tweezers, and spoon, place control brains in 5 mL of the ice-cold isolation medium in Petri dish kept on ice and immediately use for mitochondrial isolation. 5. Place each brain from the 120 min ischemia and 120 min ischemia + rotenone groups in separate wells of 24-well cell culture plate each filled with 1 mL of warm (37
C) isolation medium. 6. Transfer the cell culture plate with brains to the continuous gas flow hypoxia chamber, close the chamber and purge it with a hypoxic gas mixture. Leave the plate for 120 min at 37.0–37.5
C. Maintain the constant minimal hypoxic gas mixture flow in and out of the chamber during the experiment. 3.2 Isolation of Mitochondria from Rat Cortex and Cerebellum

All isolation procedures should be performed on ice in a cold (4
C) room (see Note 1). 1. Wash brains in 5 mL of ice-cold isolation medium in the Petri dish, separate cortex and cerebellum with the aid of scissors and tweezers, and use them further separately (see Note 1). 2. Place cortex or cerebellum in the Petri dish kept on ice and cut the tissue by scissors. 3. Take a tube with 15 mL of ice-cold isolation medium, pour little amount of it in the Petri dish with minced cortex or cerebellum, transfer the minced tissue together with 15 mL of isolation medium to a glass-Teflon homogenizer. 4. Homogenize the tissue (5 up and down strokes of the pestle) manually keeping the homogenizer in ice bath. 5. Transfer the tissue homogenate to a 50 mL centrifuge tube (see Note 2). 6. Carefully balance the centrifuge tubes in pairs with isolation medium, place tubes in the refrigerated centrifuge (4
C) each pair directly opposite one another, and spin for 5 min at 1000  g to separate large debris. 7. Decant the supernatant to new 50 mL centrifuge tubes, balance them, place in the centrifuge and spin for 10 min at 10,000  g at 4
C. 8. Discard all supernatant, turn the tubes upside down on filter paper to remove all the remaining drops. 9. Add 140–180 μL of ice-cold isolation medium to the tube with cortex mitochondria pellet—or 120–140 μL—to the tube with cerebellum mitochondria pellet and carefully re-suspend it with aid of 200 μL micropipette. Keep tubes with isolated mitochondria in ice bath during all further experimental procedures.

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10. Determine total mitochondrial protein content in each mitochondrial preparation by the modified Biuret method with BSA as standard [14]. 3.3 Measurement of Calcium Retention Capacity (CRC) in Isolated Brain Mitochondria

Ca2+-induced mPTP opening referred as CRC of isolated brain mitochondria is determined fluorimetrically using Calcium Green 5 N fluorescent dye. Measurements are performed at room temperature (25
C). 1. Prepare a fluorimeter for the fluorescence measurement (excitation at 507 nm, emission at 536 nm). 2. Take a 3 mL cuvette, add 3 mL of incubation buffer, 5 mM of succinate, and 100 nM of fluorescent dye Calcium Green 5 N (see Note 3). Mix well after each addition with a thin glass rod, close the lid and register the baseline. Calibrate the fluorescence signal by the addition of known amounts of CaCl2 (see Note 4). 3. Take a 3 mL cuvette, add 3 mL of incubation buffer, 5 mM of succinate, mitochondria (0.6 mg protein), and 100 nM of Calcium Green 5 N. Mix with a glass rod after each addition. Close the lid of the fluorimeter and register the baseline. 4. Add 1.67 μM of CaCl2 every 120 s gently mixing with a glass rod afterwards until the sudden increase in fluorescence shows spontaneous release of Ca2+ due to the mPTP opening referred as CRC (see Fig. 1).

3.4 Assessment of Mitochondrial Respiration Rate

Mitochondrial respiration rate is registered as a decrease in dissolved O2 in the respiration medium in time assessed by a Clarktype oxygen electrode (see Note 5) in a closed thermostated glass incubation chamber (see Note 6). Measurements are performed at 37
C. 1. Wash a 2 mL incubation chamber with 70% ethanol then rinse it three times with double-distilled water, remove the liquid with the aid of an aspirator. 2. Fill the incubation chamber with the respiration medium, let it equilibrate for a while then close the stopper (see Note 7). 3. Add mitochondrial respiratory chain Complex I substrates— pyruvate and malate (1 + 1 mM) and start the registration of the baseline (~2 min). 4. Add mitochondria (0.25 mg/mL protein), register the proton leak-driven respiration rate (VLeak) [3] for 3–5 min (see Fig. 2). 5. Add 2 mM of ADP, register ADP-stimulated phosphorylating respiration rate (VADP) for ~5 min. 6. Add mitochondrial respiratory chain Complex I inhibitor amytal (1.5 mM), register the respiration rate for ~2 min.

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Perfusion of Rot + Ischemia 1000 950

Control

Ischemia

900 850 800

Relative fluorescence units

750 700 650 600 550

Ca2-

500 450 400 350 300 250 200 150 100 50 0

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Fig. 1 Typical traces of calcium (Ca2+) retention capacity measurement in isolated cortex mitochondria. Rot— rotenone

Fig. 2 Typical traces of respirometric recording of cortex mitochondria. Additions: MT—mitochondria (0.25 mg protein/mL) in the presence of pyruvate/malate (1 + 1 mM); ADP—2 mM; AM—1.5 mM amobarbital; SUCC— 5 mM succinate; ATR—100 μM atractyloside. A trace indicates oxygen concentration (nmol/mL), B—oxygen flux (pmol O2/(s  mL)

7. Add mitochondrial respiratory chain Complex II substrate succinate (5 mM), register Complex II-dependent ADP-stimulated respiration rate for ~5 min.

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8. Add blocker of ADP transport to mitochondrial matrix—atractyloside (100 μM), register atractyloside-inhibited succinatedependent leak respiration rate (VAtr) for 3–4 min. 9. Calculate respiration rates as the negative time derivative of oxygen concentration and express the rates in pmol O2/s/mg mitochondrial protein (see Note 8). 3.5 Evaluation of the Activity of Complex I of Mitochondrial Respiratory Chain

Complex I activity is determined spectrophotometrically by following the kinetics of NADH oxidation at a wavelength of 340 nm Measurements are performed at room temperature (25
C). 1. Prepare submitochondrial particles (SMP): (a) Transfer 100–200 μL aliquots of isolated cortex or cerebellum mitochondria to Eppendorf test tubes (see Note 9). (b) Freeze Eppendorf test tubes with mitochondria at 20
C. (c) Thaw Eppendorf test tubes with mitochondria in the water bath at 37
C. (d) Repeat b and c procedures 3–5 times. (e) Sonicate Eppendorf test tubes with frozen/thawed mitochondria for 5 min in a refrigerated ultrasound unit. 2. Prepare a spectrophotometer for the absorption measurement at 340 nm. 3. Take a 1 mL cuvette. Add 1 mL of phosphate (25 mM) buffer supplemented with 3 mg/mL bovine serum albumin (BSA), SMP (0.125 mg protein/mL), 60 μM coenzyme Q1, 2 μg/mL antimycin A, 2 mM sodium azide. Mix with a thin glass rod, close the lid and measure the blank control. 4. Add 100 μM NADH, mix with a rod and follow NADH oxidation kinetics for 2 min. 5. Add mitochondrial respiratory chain Complex I inhibitor rotenone (10 μM), mix well and register NADH oxidation kinetics for 2 more min (see Fig. 3). 6. Calculate Complex I activity as the difference between NADH oxidation rate without and with rotenone (10 μM) and express it as nmol/min mg protein.

3.6 Evaluation of the Activity of Complex II of Mitochondrial Respiratory Chain

Complex II activity is evaluated by following the reduction of 2,6-dichlorophenolindophenol (DCPIP) spectrophotometrically according to the decrease of absorbance at 600 nm of the oxidized DCPIP. Measurements are performed at room temperature (25
C).

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Fig. 3 Typical traces of Complex I activity measurement in submitochondrial particles prepared from isolated cortex mitochondria

1. Prepare SMP (see step 1 of Subheading 3.5). 2. Prepare a spectrophotometer for the absorption measurement at 600 nm. 3. Take a 1 mL cuvette. Add 1 mL of phosphate (50 mM) buffer, 0.1 mM EDTA, 2 mM sodium azide, 10 mM succinate, SMP (0.125 mg protein/mL). Mix with a thin glass rod, close the lid, and incubate for 10 min. 4. Start the reaction by adding 1.63 mM phenazine methosulfate and 35 μM DCPIP, mix with the rod, close the lid, and follow the reduction of DCPIP for 2 min. 5. Calculate Complex II (succinate dehydrogenase (SDH)) activity (μmol/min mg protein) as the DCPIP reduction rate according to the formula: SDH activity ¼

ΔA=Δt V   103 m ε

where ΔA—absorption change; Δt—time change (min.); ε—DCPIP extinction coefficient (ε340 ¼ 19.1 mM1 cm1); V—reaction volume (mL); m—amount of mitochondrial protein (mg). 3.7 Assessment of H2O2 Generation in Isolated Brain Mitochondria

H2O2 generation in isolated mitochondria is estimated fluorimetrically with Amplex® Red reagent (10-acetyl-3,7-dihydroxyphenoxazine) and horseradish peroxidase Type IV-A. In the presence of peroxidase, the Amplex® Red reagent reacts with H2O2 in a 1:1 stoichiometry to produce the red-fluorescent oxidation product, resorufin. Measurement is performed at room temperature 25
C under constant shaking, which is stopped automatically during sample reading.

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Fig. 4 Resorufin fluorescence signal calibration by H2O2

1. Prepare fluorescence plate reader for the fluorescence measurement (excitation at 544 nm, emission at 590 nm). 2. Take black flat bottom 96 well microtiter plate, add 200 μL of respiration medium in each sample well. 3. Add substrates: pyruvate and malate (1 + 1 mM) or succinate (5 mM), or both, according to the experimental design. 4. Add rotenone (1 μM) to the wells were succinate alone is used as substrate. 5. Add 10 μM Amplex® Red and 5 U/mL horseradish peroxidase Type IV-A. 6. Add mitochondria (0.05 mg/mL). 7. Read the fluorescence signal at 0, 15 and 30 min. 8. Calibrate fluorescence signal (see Fig. 4) using 10 μM Amplex® Red, 5 U/mL horseradish peroxidase Type IV-A and known amounts (0.05 nmol, 0.13 nmol, 0.2 nmol, 0.3 nmol, 0.4 nmol) of H2O2 (see Note 10). 9. Express the rate of H2O2 generation in pmol/min/mg protein, normalized to 100% of control mitochondria.

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Notes 1. Steps 1–5 of mitochondrial isolation procedure are repeated very quickly for each experimental group and tissue type while the rest of brains are kept in isolation medium on ice. 2. Centrifuge tubes with different tissue homogenate from different experimental groups are marked and left in ice bath until all the samples are homogenized. 3. Stock solutions of all dyes should be kept in dark Eppendorf tubes. 4. Usually two additions of 1.67 μM of CaCl2 are required to cover full scale of fluorescence signal. 5. OROBOROS Oxygraph-2 k (Oroboros Instruments, Innsbruck, Austria) is the most common measuring instrument for mitochondrial functional analysis. 6. 1–2.5 mL incubation chamber is usually used to assess mitochondrial respiration rate. Teflon-coated stirrer bar is placed inside the chamber, and the chamber is mounted on an electromagnetic stirrer plate. The respiration medium is constantly stirred throughout the experiment at ~750 rpm. 7. No air bubbles should be left in the incubation chamber, the respiration medium should also fill the capillary tube in the chamber stopper through which the additions of chemicals and mitochondria are made by the aid of microsyringes. 8. Oxygen solubility was taken to be 211 nmol/O2 at 37
C [15]. 9. Mitochondria in Eppendorf tubes can be stored at 20
C for later measurements of the activities of Complex I and Complex II of mitochondrial respiratory chain. 10. Fresh 3% H2O2 stock solution should be used.

References 1. Matsumoto S, Friberg H, Ferrand-Drake M, Wieloch T (1999) Blockade of the mitochondrial permeability transition pore diminishes infarct size in the rat after transient middle cerebral artery occlusion. J Cereb Blood Flow Metab 19(7):736–741. https://doi.org/10. 1097/00004647-199907000-00002 2. Schinzel AC, Takeuchi O, Huang Z, Fisher JK, Zhou Z, Rubens J, Hetz C, Danial NN, Moskowitz MA, Korsmeyer SJ (2005) Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc Natl Acad Sci U S A 102(34):12005–12010. https://doi. org/10.1073/pnas.0505294102

3. Griffiths EJ, Halestrap AP (1993) Protection by Cyclosporin a of ischemia/reperfusioninduced damage in isolated rat hearts. J Mol Cell Cardiol 25(12):1461–1469. https://doi. org/10.1006/jmcc.1993.1162 4. Eliseev RA, Filippov G, Velos J, VanWinkle B, Goldman A, Rosier RN, Gunter TE (2007) Role of cyclophilin D in the resistance of brain mitochondria to the permeability transition. Neurobiol Aging 28(10):1532–1542. https://doi.org/10.1016/j.neurobiolaging. 2006.06.022 5. Tsuji A, Tamai I, Sakata A, Tenda Y, Terasaki T (1993) Restricted transport of cyclosporin a across the blood-brain barrier by a multidrug

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Ramune Morkuniene et al.

transporter, P-glycoprotein. Biochem Pharmacol 46(6):1096–1099. https://doi.org/10. 1016/0006-2952(93)90677-o 6. Brustovetsky N, Dubinsky JM (2000) Limitations of cyclosporin a inhibition of the permeability transition in CNS mitochondria. J Neurosci Off J Soc Neurosci 20(22): 8229–8237. https://doi.org/10.1523/ jneurosci.20-22-08229.2000 7. Gijtenbeek JM, van den Bent MJ, Vecht CJ (1999) Cyclosporine neurotoxicity: a review. J Neurol 246(5):339–346. https://doi.org/10. 1007/s004150050360 8. Sherer TB, Richardson JR, Testa CM, Seo BB, Panov AV, Yagi T, Matsuno-Yagi A, Miller GW, Greenamyre JT (2007) Mechanism of toxicity of pesticides acting at complex I: relevance to environmental etiologies of Parkinson’s disease. J Neurochem 100(6):1469–1479. https://doi.org/10.1111/j.1471-4159.2006. 04333.x 9. Barreto-Torres G, Parodi-Rulla´n R, Javadov S (2012) The role of PPARα in metformininduced attenuation of mitochondrial dysfunction in acute cardiac ischemia/reperfusion in rats. Int J Mol Sci 13(6):7694–7709. https:// doi.org/10.3390/ijms13067694 10. Chen Q, Lesnefsky EJ (2011) Blockade of electron transport during ischemia preserves bcl-2 and inhibits opening of the mitochondrial permeability transition pore. FEBS Lett 585(6):

921–926. https://doi.org/10.1016/j.febslet. 2011.02.029 11. Skemiene K, Rekuviene E, Jekabsone A, Cizas P, Morkuniene R, Borutaite V (2020) Comparison of effects of metformin, phenformin, and inhibitors of mitochondrial complex I on mitochondrial permeability transition and ischemic brain injury. Biomolecules 10(10): 1 4 0 0 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / biom10101400 12. Rekuviene E, Ivanoviene L, Borutaite V, Morkuniene R (2017) Rotenone decreases ischemia-induced injury by inhibiting mitochondrial permeability transition in mature brains. Neurosci Lett 653:45–50. https://doi. org/10.1016/j.neulet.2017.05.028 13. Rekuviene E, Ivanoviene L, Borutaite V, Morkuniene R (2017) Data on effects of rotenone on calcium retention capacity, respiration and activities of respiratory chain complexes I and II in isolated rat brain mitochondria. Data Brief 13:707–712. https://doi.org/10.1016/j.dib. 2017.06.052 14. Gornall AG, Bardawill CJ, David MM (1949) Determination of serum proteins by means of the biuret reaction. J Biol Chem 177(2): 751–766 15. Holtzman JL (1976) Calibration of the oxygen polarograph by the depletion of oxygen with hypoxanthine-xanthine oxidase-catalase. Anal Chem 48(1):229–230. https://doi.org/10. 1021/ac60365a048

Chapter 4 Assessment of Mitochondrial Complex II and III Activity in Brain Sections: A Histoenzymological Technique Rubina Roy, Rajib Paul, Pallab Bhattacharya, and Anupom Borah Abstract Mitochondrial impairment stands to be a major factor which contributes to the onset and pathogenesis of several neurodegenerative disorders, of which Alzheimer’s disease (AD), Parkinson’s disease (PD), and Huntington’s disease (HD) are among the notable ones. Extensive researches suggest the probable role of mitochondrial complex II and III dysfunction as underlying players in the pathogenesis of AD, PD, and HD. Present scenario of the world in occurrence of neurodegenerative disorders demands more research and development in this field. The development of enzyme histochemistry as an analytical technique has eased the assessment of mitochondrial complex activity at both qualitative and quantitative levels. Based on the principle of redox reactions of chromogenic substrates catalyzed by the enzymes in question, this histochemical analysis has been applied by researchers worldwide and has proved to be reliable. The present chapter hereby discusses the methods followed in performing histoenzymology of mitochondrial complex II and III activity. The chapter also puts light on the precautions which should be followed while performing histoenzymology in order to yield significant results. Key words Alzheimer’s disease, Parkinson’s disease, Huntington’s disease, Mitochondrial complex dysfunction, Succinate dehydrogenase, Coenzyme Q-cytochrome c oxidoreductase, Enzyme histochemistry

Abbreviations AD DAB HD K2HPO4 KH2PO4 MPTP Na2HPO4 NaCl

Alzheimer’s disease 3,30 -Diaminobenzidine Huntington’s disease Potassium phosphate dibasic Potassium phosphate monobasic 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine hydrochloride Sodium phosphate dibasic Sodium chloride

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-2309-1_28 Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, Corrected Publication 2022

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NaH2PO4 NBT PBS PD SDH

1

Sodium phosphate monobasic Nitro Blue Tetrazolium Sodium phosphate buffer Parkinson’s disease Succinate dehydrogenase

Introduction Due to high requirement of metabolic energy, neurons remain highly dependent on mitochondria and thereby are more susceptible to the adverse consequences of mitochondrial dysfunction. Impairment of structural and functional aspects of mitochondria is considered as a major reason behind several forms of neurodegenerative disorders [1–3]. Recent evidences suggest the major contribution of mitochondrial dysfunction at the respiratory chain complexes in the pathogenesis of neurodegenerative disorders such as AD, PD, and HD. Characterized by mild to severe impairment of memory and cognitive function, the neuropathogenesis of AD is associated with reduced functionality of complex II and III. Age-dependent decline in complex II activity has been demonstrated in mice model of AD [1]. Another study reported global deline in expression of complex III core protein in several regions of brain of AD patients, thereby directing toward a reduction in complex III activity [4]. Impaired activity of complex II and III has also been associated with the pathogenesis of PD, a neurodegenerative locomotor disorder. Spectrophotometric analysis showed a promising decline in activities of both complex II and III in brain regions of PD subjects [2]. Furthermore, reduced activity of complex II and III in MPTP induced PD model of mice was demonstrated to be exaggerated by supplementation of high cholesterol diet [5]. Studies have also correlated mitochondrial complex dysfunction with the neuropathology of Huntington’s disease (HD), a neurodegenerative disorder marked by abnormal and uncontrolled body movements, cognitive impairment, and psychiatric perturbations in individuals. Reduced functionality of complex II and III was observed in the brain of HD patients [6]. Furthermore, expression of complex II protein was found to be markedly declined in brain of rodents genetically induced with HD, while its over-expression exhibited neuroprotection against HD, indicating a major contribution of complex II dysfunction in the pathogenesis of HD [3]. The above cited evidences direct toward the role of mitochondrial complex II and III dysfunction as probable markers in neurodegenerative disorders such as AD, PD, and HD. Rise in the occurrence of neurodegenerative disorders universally urges for more developments in decoding the underlying mechanisms and come up with advanced therapeutic solutions. The present chapter hence discusses about

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the method that can be applied in the assessment of complex II and III activity in brain. One of the promising techniques applied by researchers all around the globe to assess the functionality of mitochondrial complexes is enzyme histochemistry [5, 7–10]. Enzyme histochemistry, also known as histoenzymology, has been practiced in various researches since long back and has emerged out as one of the most useful analytical techniques for evaluating the functionality of enzymes. It is applied in assessing the biochemical changes in cell brought about by enzymatic modulations as a consequence to drugs and toxins as well as endogenous factors. Enzyme histochemistry is aided by the involvement of chromogenic substrates (dyes) which get metabolized by the enzyme and thereby yields colored products whose density marks the equivalence to enzyme activity [11]. Moreover, this histochemical analysis allows the visualization of enzyme activity in individual cells present in a complex histology and that too without hindering the structural integrity of tissues [12]. Mitochondrial complex II (succinate dehydrogenase, SDH) histoenzymology is based on the principle that the enzyme SDH catalyzes the oxidation of substrate succinate and the donated electrons are received by Nitro Blue Tetrazolium (NBT). NBT is water soluble and pale yellow in color having nitro blue as the chromogenic substrate. Upon accepting electrons, the pale yellow tetrazolium gets reduced to a purple-blue colored and water insoluble compound formazan, the density of which is directly proportional to the enzymatic activity of complex II [13, 14] (Fig. 1). While the principle behind complex III (CoQ-cytochrome c oxidoreductase) histoenzymology involves the oxidation of 3,30 -Diaminobenzidine (DAB) catalyzed by enzyme cytochrome c oxidoreductase and the donated electrons are received by cytochrome c. DAB is a water soluble and colorless organic compound, which on losing electrons gets oxidized into a dark brown colored

Fig. 1 A schematic diagram illustrating the oxidoreduction of substrates by complex II (SDH)

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Fig. 2 A schematic diagram illustrating the oxidoreduction of substrates by complex III (coenzyme Q-cytochrome c oxidoreductase)

precipitate, the density of which is directly proportional to the intensity of enzymatic activity of complex III [14] (Fig. 2). The present chapter hereby discusses about the methodology involved in performing enzyme histochemistry of mitochondrial complex II and III in mice brain sections.

2

Materials

2.1 Histoenzymology of Mitochondrial Complex II (Succinate Dehydrogenase, SDH) Activity 2.1.1 Solutions and Reagents

PBS (0.1 M, pH 7.4), glycerol (10% v/v), sucrose (30% v/v), polyL-lysine, potassium phosphate buffer (0.05 M), NBT (0.03 M), sodium succinate (0.05 M). 1. Sodium Phosphate Buffer solution preparation (0.1 M, 1000 ml, pH 7.4) (see Note 1). (a) Take a little amount of distilled water in a 1000 ml measuring cylinder, add 10.987 g Na2HPO4 (i.e., 0.1 M) to it, and vortex the solution till fine dilution. (b) Add 2.711 g NaH2PO4 (i.e., 0.1 M) to above solution and vortex it till fine dilution. (c) Add 0.9% NaCl to the above made solution and vortex it till fine dilution. (d) Make the final volume of solution up to 1000 ml and adjust the pH to 7.4. (e) Pour in a clean bottle and keep it inside refrigerator. 2. Glycerol preparation (10% v/v, 100 ml):- Add 10 g glycerol (molar mass 92.09 g/mol) in 100 ml distilled water and store it in a glass bottle.

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3. Sucrose solution preparation (30% w/v, 100 ml) (see Note 1). (a) Take 70 ml of above prepared PBS in a 500 ml measuring cylinder, and add 30 g sucrose to it. (b) Vortex the solution till fine dilution. (c) Add 30 ml more PBS to the solution to make the volume up to 100 ml. (d) Pour in a clean glass bottle and keep it inside refrigerator. 4. Potassium Phosphate buffer preparation (0.05 M, 100 ml, pH 7.4) (see Note 2). (a) Take a little amount of distilled water in a 500 ml measuring cylinder. (b) Add 0.59 g KH2PO4 (i.e., 0.05 M) to the above taken distilled water and vortex it till fine dilution. (c) Add 0.115 g K2HPO4 (i.e., 0.05 M) to the above solution and vortex it till fine dilution. (d) Add more amount of water to make the final volume up to 100 ml and adjust the pH up to 7.4. 5. Reaction mixture preparation (0.1 ml) (see Note 3). (a) Take a little amount of prepared potassium phosphate buffer (0.05 M) in a 25 ml measuring cylinder. (b) Add 0.00245 g NBT (0.03 M) to above taken phosphate buffer and vortex it till fine dilution. (c) Add 0.00135 g sodium succinate (0.05 M) to above mixture and vortex it till fine dilution. (d) Add more amount of phosphate buffer to make the final volume up to 100 μl (0.1 ml). (e) Pour it in a dark or covered container and keep away from light. 2.2 Histoenzymology of Mitochondrial Complex III (CoenzymeQCytochrome c Oxidoreductase) Activity 2.2.1 Solutions and Reagents

Sodium Phosphate Buffer solution (PBS), glycerol (10% v/v), 30% w/v sucrose, poly-L-lysine, 3,30 -diaminobenzidine (DAB), cytochrome c. 1. Sodium Phosphate Buffer solution preparation (0.1 M, 1000 ml) (see Note 1). (a) Take a little amount of distilled water in a 1000 ml measuring cylinder, add 10.987 g Na2HPO4 (0.1 M) to it, and vortex the solution till fine dilution. (b) Add 2.711 g NaH2PO4 (0.1 M) to solution 1 and vortex it till fine dilution. (c) Add 0.9% NaCl to the above made solution and vortex it till fine dilution.

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(d) Make the final volume of solution up to 1000 ml and adjust the pH to 7.4. (e) Pour in a clean glass bottle and keep it inside refrigerator. 2. Glycerol preparation (10% v/v, 100 ml):- Add 10 g glycerol (molar mass 92.09 g/mol) in 100 ml distilled water and store it in a glass bottle. 3. Sucrose solution preparation (30% w/v, 100 ml) (see Note 1). (a) Take 70 ml of above prepared PBS in a 500 ml measuring cylinder, and add 30 g sucrose to it. (b) Vortex the solution till fine dilution. (c) Add 30 ml more PBS to the solution to make the volume up to 100 ml. (d) Pour in a clean glass bottle and keep it inside refrigerator. 4. Reaction mixture preparation (10 ml) (see Note 3). (a) Take a little amount of above prepared PBS (0.1 M, pH 7.4) in a 25 ml measuring cylinder. (b) Add 5.6 mg DAB to taken PBS and vortex it till fine dilution. (c) Add 2.33 mg cytochrome c to above mixture and vortex it till fine dilution. (d) Add 500 mg sucrose to above mixture and vortex it till fine dilution. (e) Lastly, add more amount of PBS to make the final volume up to 10 ml. (f) Pour it in a dark or covered container and keep away from light.

3

Method

3.1 Histoenzymology of Mitochondrial Complex II (Succinate Dehydrogenase, SDH) Activity

1. Anesthetize the animals and perfuse transcardially with 0.1 M PBS (pH 7.4) and 10% v/v glycerol. 2. Post-perfusion, dissect out the brain from calvarium and preserve it in 30% w/v sucrose solution for whole night at 4 c temperature for cryopreservation. 3. Following cryopreservation period, take out the brain from sucrose solution and prepare it for sectioning in cryomicrotome. 4. Collect serial sections of brain passing through specific regions of interest with desirable thickness on poly-L-lysine coated slides, and wash with PBS (0.1 M, pH 7.4) for 10 min (see Notes 4 and 5).

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5. Following washing, incubate the sections in prepared reaction mixture in dark at 37  C for 30 min (see Notes 6 and 7). 6. Post-incubation, wash the sections with distilled water and mount in glycerol. 7. Gently put coverslip over the sections and immediately take fine quality photographs under bright field illumination (see Notes 8 and 9). 8. Calculate the optical density of the sections on ImageJ software (Fiji version) to evaluate complex II activity [10]. 3.2 Histoenzymology of Mitochondrial Complex III (CoenzymeQCytochrome c Oxidoreductase) Activity

1. Anaesthetize the animals and perfuse transcardially with 0.1 M PBS (pH 7.4) and 10% v/v glycerol. 2. Post-perfusion, dissect out the brain from calvarium and preserve it in 30% w/v sucrose solution for whole night at 4  C for cryopreservation. 3. Post-cryopreservation, take out the brain from sucrose solution and prepare it for sectioning in cryomicrotome. 4. Collect serial sections of brain passing through specific regions of interest with desirable thickness on poly-L-lysine coated slides and rinse the sections in PBS (0.1 M, pH 7.4) for three times (see Notes 4 and 5). 5. Incubate the sections in prepared reaction mixture in dark at 37 c for 1 h (see Notes 6 and 7). 6. Post-incubation, wash the sections in PBS (0.1 M, pH 7.4) and mount in glycerol. 7. Gently put coverslip over the sections and immediately take fine quality photographs under bright field illumination (see Notes 8 and 9). 8. Calculate the optical density of the sections on ImageJ software (Fiji version) to evaluate complex III activity [10].

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Notes 1. PBS and sucrose solutions prepared can be stored in refrigerator for a month. 2. Potassium phosphate buffer kept in a dark or properly foiled container and stored inside refrigerator can be used for a month, however, freshly prepared is preferable. 3. Always prepare the reaction mixture right before use and stock it in a dark or properly foiled container. 4. Pre-coat the slides with poly-L-lysine few hours before use and allow it to dry for better coating.

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5. Keep the section loaded slides in a moist chamber so as to avoid dryness and tissue damage. 6. Perform the incubation of sections in reaction mixture in dark so as to avoid interference from light (as light degrades the chromogenic substrate). 7. Never forget to make a water-resistant boundary encircling the sections on the slide with the help of a water-proof marker pen, before incubation. This will restrict the reaction mixture from getting drained out. 8. Put the coverslip gently upon slides and take care of bubble formation. 9. Place the slides under microscope carefully such that glycerol does not come in contact with the lenses.

Acknowledgments We sincerely acknowledge the support received from Department of Biotechnology, Government of India (Sanction Order No. BT/PR17127/NER/95/453/2015, dated January 13, 2017). References 1. Emmerzaal TL, Rodenburg RJ, Tanila H et al (2018) Age-dependent decrease of mitochondrial complex II activity in a familial mouse model for Alzheimer’s disease. J Alzheimers Dis 66:75–82 2. Foti SC, Hargreaves I, Carrington S et al (2019) Cerebral mitochondrial electron transport chain dysfunction in multiple system atrophy and Parkinson’s disease. Sci Rep 9:1–12 3. Damiano M, Diguet E, Malgorn C et al (2013) A role of mitochondrial complex II defects in genetic models of Huntington’s disease expressing N-terminal fragments of mutant huntingtin. Hum Mol Genet 22:3869–3882 4. Kim SH, Vlkolinsky R, Cairns N, Lubec G (2000) Decreased levels of complex III core protein 1 and complex V β chain in brains from patients with Alzheimer’s disease and down syndrome. Cell Mol Life Sci 57:1810– 1816 5. Paul R, Choudhury A, Kumar S et al (2017) Cholesterol contributes to dopamine- neuronal loss in MPTP mouse model of Parkinson’s disease : involvement of mitochondrial dysfunctions and oxidative stress. PLoS One 12: e0171285

6. Browne SE, Bowling AC, Macgarvey U et al (1997) Oxidative damage and metabolic dysfunction in Huntington’s disease : selective vulnerability of the basal ganglia. Ann Neurol 41:646–653 7. Govindaiah, Shankaranarayana Rao BS, Ramamohan Y et al (2000) Cytochrome oxidase activity in rat retinal ganglion cells during postnatal development. Dev Brain Res 124:117– 120 8. Pandey M, Varghese M, Sindhu KM et al (2008) Mitochondrial NAD+-linked state 3 respiration and complex-I activity are compromised in the cerebral cortex of 3-nitropropionic acid-induced rat model of Huntington’s disease. J Neurochem 104:420– 434 9. Paul R, Borah A (2017) Global loss of acetylcholinesterase activity with mitochondrial complexes inhibition and inflammation in brain of hypercholesterolemic mice. Sci Rep 7: 1–13 10. Mazumder MK, Paul R, Bhattacharya P, Borah A (2019) Neurological sequel of chronic kidney disease: from diminished acetylcholinesterase activity to mitochondrial dysfunctions,

Histoenzymology of Mitochondrial Complexes in Brain oxidative stress and inflammation in mice brain. Sci Rep 9:1–22 11. Mazer MA, Kovacs JA, Swan JC et al (1987) Histoenzymological study of selected dehydrogenase enzymes in pneumocystis carinii. Infect Immun 55:727–730 12. Simard M, Mourier A, Greaves LC et al (2018) A novel histochemistry assay to assess and quantify focal cytochrome c oxidase deficiency. J Pathol 245:311–323

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13. Brouillet E, Guyot M, Mittoux V et al (1998) Partial inhibition of brain succinate dehydrogenase by 3-nitropropionic acid is sufficient to initiate striatal degeneration in rat. J Neurochem 70:794–805 14. Do¨lle C, Bindoff LA, Tzoulis C (2018) 3,3’Diaminobenzidine staining interferes with PCR-based DNA analysis. Sci Rep 8:1–8

Chapter 5 Measurement of Mitochondrial (Dys)Function in Cellular Systems Using Electron Paramagnetic Resonance (EPR): Oxygen Consumption Rate and Superoxide Production Donatienne d’Hose and Bernard Gallez Abstract The oxygen consumption rate (OCR) and superoxide production are crucial when assessing mitochondrial function and/or dysfunction. EPR spectroscopy allows the measurement of both components either independently or simultaneously in a same cellular or mitochondrial preparation. OCR determination using EPR oximetry is based on the change in EPR linewidth of a paramagnetic oxygen sensing probe (a perdeuterated nitroxide) in the presence of oxygen consuming cells in a closed system. Superoxide production can be monitored by the oxidation of cyclic hydroxylamines into nitroxides. The contribution of superoxide to the nitroxide formation is deduced from experiments in the presence and in the absence of SOD and PEG-SOD as appropriate controls. Key words EPR, ETC, Mitochondrial function, Oximetry, Oxygen consumption rate (OCR), Superoxide, Nitroxide, Hydroxylamine

1

Introduction Mitochondria have a pivotal role in cellular energetics including oxidative phosphorylation to produce cellular ATP, but they also have important roles in apoptosis and programmed cell death, and in ROS production. Mitochondrial dysfunction may lead to pathological processes such as diabetes, cardiovascular diseases, and neurodegenerative disorders. Over the years, several mitochondrial modulators have emerged, notably in the field of anticancer therapies targeting cellular oxygenation and mitochondrial metabolism. Their impact on the oxygen consumption rate (OCR) and bioenergetics of this organelle is nowadays well studied but little has been investigated on their potential underlying incidence on reactive oxygen species (ROS) production and oxidative damage occurring during those treatments. When the balance between antioxidant systems and cellular ROS is disrupted, overpowering oxidative

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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stress may lead to the oxidation of macromolecules like DNA, proteins and lipids, consequently inducing cellular damage and defects in cell signaling [1]. More specifically, superoxide may trigger cell death when produced in large excess but it was also suggested to be a key driver promoting cancer cell migration and metastasis when produced at more moderate levels [2]. Therefore, the dual dose-dependent effects on oxygen consumption rate (OCR) and superoxide production are crucial to consider when assessing mitochondrial function and/or dysfunction. Electron paramagnetic resonance (EPR) is a method of choice for assessing the oxygen consumption and the superoxide production. The OCR determination is based on the change in EPR linewidth of a paramagnetic oxygen sensing probe (here, a nitroxide) in the presence of oxygen consuming cells in a closed system [3– 5]. This method has been successfully applied to characterize the effect of inhibitors of the electron transport chain in cancer cells [3, 6–11]. In addition, EPR allows the detection of superoxide produced by the oxidation of cyclic hydroxylamines into nitroxides [12–15]. Because their oxidation into paramagnetic nitroxide can be due to several redox reactions, the contribution of superoxide to the nitroxide formation has to be deduced from experiments carried out in the presence and in the absence of SOD or PEG-SOD (pegylated-superoxide dismutase) as appropriate controls [14]. As described in this paper, both parameters may be measured independently. Recently, our lab also developed an integrated toolbox enabling the simultaneous analysis of the OCR and superoxide production on a single mitochondrial preparation [16]. For the purpose, there is a need to measure the variation of the EPR linewidth of one nitroxide, on the one hand, and the formation of another nitroxide, on the other hand. The separation of the EPR signals coming from both nitroxides can be done by using different isotopically-labeled probes: 15N-PDT (4-oxo-2,2,6,6-tetramethylpiperidine-d16-15N1-oxyl) is the nitroxide used for OCR measurement and 14 N-CMH (1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine) is the cyclic hydroxylamine that will be transformed into a nitroxide after reaction with the superoxide. The EPR signal coming from both nitroxides are not superimposable as 15N-PDT gives a doublet while 14N-CMH gives a triplet (Fig. 1). Therefore, the variation of the linewidth of 15N-PDT (linked to OCR) and the formation of 14N-CM (linked to superoxide production) can be recorded simultaneously on a single cellular or mitochondrial preparation [16]. l

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Fig. 1 Representative global EPR spectrum of 15N-PDT and 14N-CMH in PBS buffer injected into two separate capillaries, and gathered into a same quartz tube. Final concentrations of the probes are 100 μM and 0.5 mM, respectively

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Materials

2.1 EPR Spectrometer

2.2

EPR Oximetry

All EPR experiments were performed using a Bruker EMX-Plus spectrometer operating in X band (9.85 GHz) and equipped with a PremiumX ultra low noise microwave bridge and a SHQ high sensitivity resonator (see Note 1). The EPR cavity was heated at 310 K with air during all experiments. OCR and superoxide measurements can be performed on whole cells as well as on isolated mitochondria without changing material nor method. Specific concentrations for both models are informed for each assay. Skip the mitochondrial isolation part if using a whole cell model. 1. Disposable microhematocrit capillaries in soda-lime glass (75 mm/75 μL). 2. Sealing gum for capillaries (see Note 2). 3. Phosphate buffered saline (PBS) from Gibco, pH 7.4, without calcium chloride nor magnesium chloride. 4. 20% Dextran from leuconostoc mesenteroides (average MW 60000–76000) solution diluted in PBS (see Note 3). Stock solution can be stored at 4  C during a month. 5.

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N-PDT (4-oxo-2,2,6,6-tetramethylpiperidined16-15N-1oxyl) used as oxygen sensor (see Note 4). The required stock solution is 2 mM diluted into PBS. Store at 4  C for short term use. Other aliquots should be stored at 80  C until needed.

6. Biological system: depending on the type of experiment wanted: Stock solution of whole cells (around 5.106 cells/mL culture media) (see Note 5) or suspension of isolated mitochondria.

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2.3 Superoxide Measurements

1. Gas-permeable polytetrafluoroethylene (PTFE) tubing (ZEUS) with inside diameter 0.025 in. and wall thickness 0.002 in., cut into 12 cm sections. 2. 1 mL syringes with 23G needles. 3. PBS (Gibco), pH 7.4, without calcium chloride nor magnesium chloride. 4. Parafilm. 5. Diethylenetriaminepentaacetic acid (DTPA): 100 mM stock solution in PBS. Store at room temperature. 6. Polyethylene glycol-superoxide dismutase (PEG-SOD): 4000 U/mL stock solution in PBS. Store at 20  C. 7. Diamagnetic hydroxylamine for oxidation measurements: 1-Hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine. HCl (CMH hydrochloride) used on mitochondria or (2-(2,2,6,6-Tetramethylpiperidin-1-oxyl-4-ylamino)-2oxoethyl) triphenylphosphonium chloride (MitoTempoH) for whole cell experiments. Stock solution are 10 mM and 1 mM, respectively, in PBS. Both probes are to be stored at 20  C and flushed with argon at every use to avoid probe oxidation. 8. Biological system: depending on the type of experiment wanted: Stock solution of whole cells (around 20.106cells/ mL culture media) or suspension of isolated mitochondria.

2.4 Mitochondrial Isolation

1. Cell line of interest, cultured in its recommended medium: about 40.106cells required resuspended in 1 mL of PBS (see Note 6). 2. Motor-driven Glass/Teflon potter homogenizer. 3. 0.1 M Tris/MOPS: Dissolve 1.21 g Tris in 40 mL distillated water, then adjust pH to 7.4 with MOPS powder. Bring to 50 mL and store at 4  C. 4. 1 M EGTA/Tris: Dissolve 3.81 g EGTA in 40 mL distillated water, then adjust pH to 7.4 with Tris powder. Bring to 50 mL and store at 4  C. 5. 1 M sucrose: Dissolve 34.2 g Sucrose in 100 mL distilled water and separate in 5 aliquots. Store at 20  C. 6. Mitochondrial isolation buffer (100 mL to aliquot): Mix 10 mL of Tris/MOPS (0.1 M) and 1 mL of EGTA/Tris (1 M) to 20 mL of sucrose (1 M). Bring the mixture to 100 mL with distilled water and adjust pH to 7.4. Store at 20  C.

2.5

Mitotoolbox

The Mitotoolbox combines OCR and superoxide measurements at the same time, meaning that all material listed for OCR and superoxide are also required for this experiment. Solutions and buffers

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are kept on ice during the mitochondrial isolation procedure. Listed substrate supply includes NADH- and FADH2-linked substrates to support state 3 respiration. 1. Isolated mitochondria (see “mitochondrial isolation” in Methods). 2. 100 mM EGTA: Dissolve 76 mg in 20 mL of water. 3. Mitochondrial Assay buffer (MAS (2): This buffer is two times concentrated: Combine 12 g Sucrose, 20 g Mannitol, 68 mg potassium phosphate monobasic (KH2PO4), 0.5 g MgCl2 hexahydrate, 23.8 mg HEPES, 1 g FA-free BSA and 5 mL of EGTA stock solution (100 mM) in 250 mL of water. Adjust pH to 7.4 and aliquot. Store at 20  C. 80  C.

4. 100 mM Pyruvate: dilute in PBS. Store at 5. 500 mM Succinate: dilute in PBS. Store at 6. 500 mM Malate: dilute in PBS. Store at

80  C.

80  C.

7. 100 mM ADP: Mix 425 mg ADP with 1.2 mL of PBS (ADP does not dissolve at this point). Neutralize with 5 M KOH (approx. 450 μL). ADP will dissolve after addition of KOH. Adjust pH to 7.4 and bring to 10 mL with PBS. Store at 80  C. 8. Substrate Mix: Combine 300 μL of MAS (2) with 216 μL PBS, 60 μL pyruvate (100 mM), 6 μL succinate (500 mM), 6 μL Malate (500 mM), and 12 μL ADP (100 mM). This final volume is enough for six combined measurements of OCR and Superoxide.

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Methods

3.1 EPR Oximetry on Whole Cell Model

Keep the cell stock solution (lid open) in a 37  C water bath during the experiments. 1. Turn on the X band EPR, heat the cavity at 310 K using continuous nitrogen flow (400 L/h) (see Note 7), and open Bruker Xenon Spin fit program. 2. Tune the EPR cavity at 19 dB attenuation (2.518 mW) with a sealed hematocrit capillary filled with PBS transferred in a quartz tube (see Note 8). 3. Set experimental parameters for acquisition: microwave power: 2.518 mW; modulation frequency: 100 kHz; modulation amplitude: 0.005 mT; Center field: 335mT; Sweep width: 1.5 mT; Sweep time:15 s. 4. Combine 60 μL of cell suspension to 40 μL of Dextran 20% solution.

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Fig. 2 (a): 15N-PDT spectra at 0.1% (red) and 21% (blue) of oxygen mixed with nitrogen. (b): 15N-PDT calibration curve linking the probe’s linewidth (Gauss) to oxygen level (%) present into the sealed hematocrit capillary. Both spectra correspond to the high-field line

5. Add 4 μL of 15N-PDT stock solution to the previous mixture and mix well. 6. Transfer mix into hematocrit capillary and seal with gum. Slip capillary carefully into a quartz tube and transfer it to the EPR cavity. 7. Tune the EPR cavity and wait 3 min after the probe was mixed before launching an automated “2D-field-Delay” measurement counting 15 points with a time delay of 60,000 ms. 8. Save file and proceed to data analysis after all experiments are done: switch to processing mode and load your file. Select “Peak picking” and define the region containing the peak on all slices with “region qualifier.” Select pick > Peak and through > Dist Algorithm > All slices > Pick > Report distances. The final file can be saved as ASCII file to extract linewidth data at each point. 9. Correlate 15N-PDT linewidth with the % of oxygen with a calibration curve (see Note 9 and Fig. 2). 10. OCR corresponds to the slope of oxygen level during time. 3.2 Superoxide Measurement on Whole Cell Model

Keep the cell stock solution (lid open) in a 37  C water bath during the experiments. 1. Turn on the X band EPR, heat the cavity at 310 K using continuous air flow (400 L/h) and open Bruker Xenon Spin fit program. 2. Tune the EPR cavity at 10 dB attenuation (20 mW) with a 12 cm long PTFE tube folded in 4 filled with PBS, transferred in an open quartz tube (see Note 10). Seal the end of the quartz tube with a patch of parafilm to avoid a potential leak into the EPR cavity. Set experimental parameters for acquisition:

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microwave power: 20 mW; modulation frequency: 100 kHz; modulation amplitude: 0.1 mT; Center field: 336.5mT; Sweep width: 1.5 mT; Sweep time: 30.48 s. 3. For control condition, combine 37 μL of cell suspension to 0.5 μL od DTPA (100 mM), 5 μL of PBS and end by mixing 7.5 μL of MitoTempoH (1 mM). To measure superoxide contribution, make another measurement using the same conditions but replacing 2.5 μL PBS by adding 2.5 μL of PEG-SOD (4000 U/mL). Let incubate 10 min before adding the MitoTempoH. It is mandatory to flush MitoTempoH stock solution with argon before and during pipetting to avoid probe oxidation. 4. Transfer mix into PTFA tube cutting using the needle. Fold in 4 and insert into the open quartz tube. Seal quartz tube with a patch of parafilm and transfer it to the EPR cavity. 5. Tune the EPR cavity and wait 3 min after the probe was mixed before launching an automated “2D-field-Delay” measurement counting 11 points with a time delay of 40,000 ms. 6. Save file and proceed to data analysis after all experiments are done: switch to processing mode and load your file (Fig. 3). Select “Integration & derivative” and define the region containing the peak in all slices. Select “Double Integration” and copy results to primary. Then, switch to time domain > Store & return > Structure > Projection > Roof. The final file can be saved as ASCII file to extract double integration (DI) values for each timepoint (Fig. 4). 7. Subtract point 1 DI from point 11 DI for each condition. Superoxide contribution is measured by subtracting mean PEG-SOD DI to mean control DI.

Fig. 3 Representative example of 14N-CMH oxidation spectra after 1 and 10 min on whole cell model. These spectra are similar to those obtained with MitoTempoH probe. Both spectra correspond to the mid-field line

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Fig. 4 Representative example of MitoTempoH DI spectra during time on isolated mitochondria. Same kind of spectra are obtained when using a whole cell model as well. Blue dots represent control condition and red dots represent PEG-SOD condition. Superoxide contribution is measured by subtracting the PEG-SOD DI to the CTR DI after 15 min 3.3 Mitochondrial Isolation

Mitochondrial isolation buffer (MIB) should be toughed in advance and stored on ice. All steps of mitochondrial isolation are performed on ice or at 4  C to avoid damage by proteases and phospholipases. The potter homogenizer is precooled on ice before use. This protocol has been adapted from [17]. 1. Harvest cultured cell line of interest to get around 40.106 cells (or more if needed, see Notes 6 and 11). 2. Resuspend cells in 1 mL PBS and centrifuge cells at 600  g at 4  C during 10 min. 3. Remove the supernatant and resuspend the cells in 3 mL of ice-cold MIB. 4. Transfer the cell mixture into the precooled potter and blend with the motor-driven pestle at 1600 rpm making 35 strokes. To maintain cooling temperature during homogenization, the entire device is placed in a box or beaker filled with ice (see Note 12). 5. Transfer the homogenate to a 15 mL falcon tube and centrifuge at 600  g at 4  C during 10 min. 6. Collect the supernatant and centrifuge at 7000  g at 4  C during 10 min. 7. Remove the supernatant and wash the pellet with 200 μL of MIB before transferring the homogenate to a 1.5 mL Eppendorf. 8. Centrifuge the suspension at 7000  g at 4  C during 10 min. 9. Discard the supernatant and gently homogenize the mitochondria pellet with the little remaining supernatant left, avoiding bubble formation. 10. Store on ice (see Note 13).

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3.4 Mitotoolbox: OCR and Superoxide Simultaneous Measurement

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The Mitotoolbox combines oximetry and superoxide measurements at the same time on the same mitochondrial isolate, meaning that all material listed for EPR oximetry and superoxide measurements are also required to perform the experiment. Of note, this simultaneous measurement of mitochondrial function can also be measured on a whole cell model using the same EPR device as well as EPR parameters with the cell stock solutions given in the Material section “EPR oximetry” and “superoxide measurements.” 1. Store mitochondrial isolate and substrate mix on ice. 2. Respiration mix (for six independent measurements): combine 275 μL substrates, 137.5 μL of MAS (2), 132 μL dextran 20%, and 5.5 μL DTPA. Mix well. 3. Superoxide mix (for six independent measurements): combine 125 μL substrates, 62.5 μL MAS (2), 60 μL PBS, and 2.5 μL DTPA. Mix well. 4. Turn on the X band EPR, heat the cavity at 310 K using continuous air flow (400 L/h) and open Bruker Xenon Spin fit program. 5. Tune the EPR cavity at 19 dB attenuation (2.518 mW) with a 12 cm long PTFE tube folded in 4 filled with PBS and a sealed hematocrit capillary filled with PBS (see Note 14 and Fig. 5). 6. Create and save a separate OCR parameters file on Xenon: microwave power: 2.518 mW; modulation frequency: 100 kHz; modulation amplitude: 0.005 mT; Receiver gain: 45 dB; Center field:334 mT; Sweep width: 1.5 mT; Sweeptime: 25 s. 7. Create and save a separate superoxide parameters file on Xenon: microwave power: 2.518 mW; modulation frequency: 100 kHz; modulation amplitude: 0.1 mT; Receiver gain: 60 dB; Center field: 334.4 mT; Sweep width: 1.5 mT; Sweeptime: 25 s. 8. Final OCR solution for EPR: combine 85 μL of respiration mix with 7 μL PBS, 3 μL of isolated mitochondria and add 5 μL of 15 N-PDT (2 mM) at the last minute. 9. Final superoxide solution for EPR: combine 32 μL superoxide mix with 3 μL PBS, 3 μL isolated mitochondria and add 2 μL of 14 N-CMH (10 mM) at the last minute. 10.

15

N-PDT and 14N-CMH should be inserted to their mix at the same time. Hematocrit capillary and PTFA tubing are filled with their corresponding final solution, and put together into the open quartz tube. Transfer the device into the EPR cavity.

11. 3 min after probes were added to their final mix, launch first superoxide acquisition with the corresponding parameters and

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Fig. 5 Schematic representation of the double capillary device used to measure OCR and superoxide simultaneously

save file. Launch OCR acquisition every even number of minutes and superoxide on odd number of minutes after probe addition. Save files on Xenon for data analysis. 12. To measure superoxide contribution, add 5 μL of PEG-SOD to final OCR solution and 2 μL PEG-SOD to the final superoxide solution (adding less PBS in consequence). 13. Data analysis are performed the same way as stated in section “EPR oximetry” and section “superoxide measurement” on whole cell model. 14. Normalization by mitochondrial protein quantification is recommended (see Note 15).

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Notes 1. Any EPR spectrometer can be used as long as it is capable of being equilibrated at 310 K with continuous gas flow. 2. Gum can contain paramagnetic manganese visible with EPR when inserted into the cavity’s resonator. To avoid nonspecific spectra appearing during analysis, make sure the sealed part of the hematocrit capillary remains outside the resonator. 3. Dextran solution is used to keep cells or isolated mitochondria in suspension in the capillary during the whole measurement. Do not use dextran with a lower MW than stated, otherwise cells tend to sedimentate and EPR measurements will not be accurate. 4.

15

N-PDT (perdeuterated nitroxide) is a very sensitive probe to measure oxygen levels in a medium using EPR technology. Its linewidth is proportional to the oxygen level present into the capillary. A calibration linking the probe linewidth broadening to oxygen level should therefore first be performed in order to measure OCR during an experiment with cells or mitochondrial suspension.

5. For whole cells experiments, a stock solution of 5.106 cells/mL should be sufficient to measure a suitable oxygen consumption. However, depending on the cell type, this cell concentration can be fine-tuned beforehand by performing dose-response tests to get the best oxygen consumption curve (some cells are more rapidly consuming oxygen than others). This new stock concentration should therefore be used for all other oximetry experiments using this cell line. 6. Depending on the cell line, this initial cell number could be increased to have a better yield and perform more measurements from the same isolated batch of mitochondria. Typically, at the end of the process, enough mitochondria are collected (40 μL with a mean of 25 mg of protein/mL) to make four conditions with simultaneous measurements of OCR and superoxide, using 3T3 fibroblasts. 7. In this case, EPR cavity can also be heated with air but nitrogen is less expensive. The capillary being sealed off, the type of gas used to heat the cavity does not matter because it will not affect the sample. 8. The tuning of the EPR cavity is performed at 19 dB attenuation (2.518 mW) with the same device used for the actual oximetry experiments, namely a sealed hematocrit capillary filled with PBS and 15N-PDT (oxygen sensor), inserted into a quartz tube that is then transferred to the EPR cavity. Mixing the oxygen sensor at this stage is not mandatory but recommended in

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order to find the exact magnetic field value where the 15N-PDT spectrum is appearing. 9. The calibration to link 15N-PDT linewidth with the level of oxygen in a capillary is performed using an EPR spectrometer connected to an Aalborg gas mixer and an oxygen analyzer (Servomex OA540). Calibration mixture contains PBS and Dextran (20%) at a final ratio 1:1 mixed with 4 μL of 15 N-PDT (2 mM). 15N-PDT linewidth is measured at different % of oxygen (7pts between 0 and 21%) mixed with nitrogen. Wait 40 s before each measurement for the oximeter to equilibrate. The obtained curve equation is then used to measure % of oxygen during experiments on biological systems. 10. To fill the PTFA tubing, insert needle into a tube segment and suck up the PBS or final cell solution until the segment is entirely filled. Fold in 4 and insert the tube at the open end of the quartz tube. 11. Do not treat cells with any agents before harvesting. Mitochondrial isolation is performed on control/non-treated cells and treatment of any kind should be added in final OCR or final superoxide solutions as a new condition to analyze its impact on mitochondrial function. 12. Homogenization by a motor-driven Glass/Teflon potter tends to heat up the cell mixture. To avoid damaging activation of proteases and phosphatases and maintain mitochondrial integrity, keeping the potter on ice is essential. 13. Isolated mitochondria should be used within 1–3 h for better responses. 14. The two tubes are inserted next to each other in the open quartz tube. 15. Protein quantification can be performed using BCA protein Assay. Mitochondrial isolates are diluted 1:20 and protein determination can be assessed following manufacturer’s instructions.

Acknowledgments This research has been supported by the Foundation against Cancer (2016-087) and the Fonds National de la Recherche Scientifique FNRS (7652717F and J008220F). References 1. Purohit V, Simeone DM, Lyssiotis CA (2019) Metabolic regulation of redox balance in cancer. Cancers (Basel) 11(7):955. https://doi. org/10.3390/cancers11070955

2. Porporato PE, Payen VL, Perez-Escuredo J, De Saedeleer CJ, Danhier P, Copetti T, Dhup S, Tardy M, Vazeille T, Bouzin C, Feron O, Michiels C, Gallez B, Sonveaux P

EPR Assays for Oxygen Consumption and Superoxide Production (2014) A mitochondrial switch promotes tumor metastasis. Cell Rep 8(3):754–766. https://doi.org/10.1016/j.celrep.2014. 06.043 3. Jordan BF, Gregoire V, Demeure RJ, Sonveaux P, Feron O, O’Hara J, Vanhulle VP, Delzenne N, Gallez B (2002) Insulin increases the sensitivity of tumors to irradiation: involvement of an increase in tumor oxygenation mediated by a nitric oxide-dependent decrease of the tumor cells oxygen consumption. Cancer Res 62(12):3555–3561 4. Diepart C, Verrax J, Calderon PB, Feron O, Jordan BF, Gallez B (2010) Comparison of methods for measuring oxygen consumption in tumor cells in vitro. Anal Biochem 396(2): 250–256. https://doi.org/10.1016/j.ab. 2009.09.029 5. Danhier P, Copetti T, De Preter G, Leveque P, Feron O, Jordan BF, Sonveaux P, Gallez B (2013) Influence of cell detachment on the respiration rate of tumor and endothelial cells. PLoS One 8(1):e53324. https://doi.org/10. 1371/journal.pone.0053324 6. Jordan BF, Sonveaux P, Feron O, Gregoire V, Beghein N, Dessy C, Gallez B (2004) Nitric oxide as a radiosensitizer: evidence for an intrinsic role in addition to its effect on oxygen delivery and consumption. Int J Cancer 109(5):768–773. https://doi.org/10.1002/ ijc.20046 7. Crokart N, Jordan BF, Baudelet C, Ansiaux R, Sonveaux P, Gregoire V, Beghein N, DeWever J, Bouzin C, Feron O, Gallez B (2005) Early reoxygenation in tumors after irradiation: determining factors and consequences for radiotherapy regimens using daily multiple fractions. Int J Radiat Oncol Biol Phys 63(3):901–910. https://doi.org/10.1016/j. ijrobp.2005.02.038 8. Crokart N, Jordan BF, Baudelet C, Cron GO, Hotton J, Radermacher K, Gregoire V, Beghein N, Martinive P, Bouzin C, Feron O, Gallez B (2007) Glucocorticoids modulate tumor radiation response through a decrease in tumor oxygen consumption. Clin Cancer Res 13(2 Pt 1):630–635. https://doi.org/10. 1158/1078-0432.CCR-06-0802 9. Ansiaux R, Baudelet C, Jordan BF, Crokart N, Martinive P, DeWever J, Gregoire V, Feron O, Gallez B (2006) Mechanism of reoxygenation after antiangiogenic therapy using SU5416 and its importance for guiding combined antitumor therapy. Cancer Res 66(19):9698–9704.

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https://doi.org/10.1158/0008-5472.CAN06-1854 10. Diepart C, Karroum O, Magat J, Feron O, Verrax J, Calderon PB, Gregoire V, Leveque P, Stockis J, Dauguet N, Jordan BF, Gallez B (2012) Arsenic trioxide treatment decreases the oxygen consumption rate of tumor cells and radiosensitizes solid tumors. Cancer Res 72(2):482–490. https://doi.org/ 10.1158/0008-5472.CAN-11-1755 11. De Preter G, Deriemaeker C, Danhier P, Brisson L, Cao Pham TT, Gregoire V, Jordan BF, Sonveaux P, Gallez B (2016) A fast hydrogen sulfide-releasing donor increases the tumor response to radiotherapy. Mol Cancer Ther 15(1):154–161. https://doi.org/10.1158/ 1535-7163.MCT-15-0691-T 12. Dikalov SI, Kirilyuk IA, Voinov M, Grigor’ev IA (2011) EPR detection of cellular and mitochondrial superoxide using cyclic hydroxylamines. Free Radic Res 45(4):417–430. https://doi.org/10.3109/10715762.2010. 540242 13. Dikalov SI, Harrison DG (2014) Methods for detection of mitochondrial and cellular reactive oxygen species. Antioxid Redox Signal 20(2): 372–382. https://doi.org/10.1089/ars. 2012.4886 14. Scheinok S, Leveque P, Sonveaux P, Driesschaert B, Gallez B (2018) Comparison of different methods for measuring the superoxide radical by EPR spectroscopy in buffer, cell lysates and cells. Free Radic Res 52(10): 1182–1196. https://doi.org/10.1080/ 10715762.2018.1541321 15. Scheinok S, Capeloa T, Porporato PE, Sonveaux P, Gallez B (2020) An EPR study using cyclic hydroxylamines to assess the level of mitochondrial ROS in superinvasive cancer cells. Cell Biochem Biophys 78(3):249–254. https://doi.org/10.1007/s12013-02000921-6 16. d’Hose D, Danhier P, Northshield H, Isenborghs P, Jordan BF, Gallez B (2021) A versatile EPR toolbox for the simultaneous measurement of oxygen consumption and superoxide production. Redox Biol 40: 101852. https://doi.org/10.1016/j.redox. 2020.101852 17. Frezza C, Cipolat S, Scorrano L (2007) Organelle isolation: functional mitochondria from mouse liver, muscle and cultured fibroblasts. Nat Protoc 2(2):287–295. https://doi.org/ 10.1038/nprot.2006.478

Chapter 6 Mitochondrial Calcium Handling in Isolated Mitochondria from a Guinea Pig Heart Jyotsna Mishra and Amadou K. S. Camara Abstract Mitochondrial calcium (Ca2+) plays a key role in regulating normal cardiac function. A physiological increase in mitochondrial matrix calcium [Ca2+]m drives mitochondrial ATP production to meet the high-energy demands during excitation–contraction coupling. However, a pathological increase in [Ca2+]m leads to increased oxidative stress, impaired bioenergetics, and the opening of mitochondrial permeability transition pore (mPTP), a hallmark of the failing heart. Therefore, a better understanding of the [Ca2+]m handling and its role in heart function and dysfunction is of great importance. Here, we describe a detailed protocol for measuring mitochondrial Ca2+ handling in the isolated functionally intact mitochondria from cardiac tissue of the guinea pig. Key words Mitochondrial calcium handling, Mitochondrial bioenergetics, Mitochondrial oxygen consumption, Calcium retention capacity, Mitochondrial membrane potential

1

Introduction The heart is one of the most metabolically active organs that require a continuous and rapid supply of energy to sustain its normal function. The high-energy demand of the beating heart is largely produced in cardiomyocytes by mitochondrial oxidative metabolism. Consequently, cardiac tissue has a high abundance of mitochondria (~35–40% of cardiomyocyte volume) [1], to ensure a highly efficient energy production (> 90% of the ATP) [2] to support excitation–contraction coupling, and to maintain ionic constancy. Thus, beyond their role as the main energy source of the myocardium, mitochondria provide important calcium buffering machinery and play a key role in the regulating intracellular calcium homeostasis. An increase in mitochondrial matrix calcium activates oxidative phosphorylation and ATP generation, in part, by stimulating, key enzymes of the tri-carboxylic acid cycle [3] to fuel the contraction. Mitochondria also serve other cellular functions,

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such as, generating and regulating reactive oxygen species (ROS), apoptotic activation and contribution to cell survival and death. Given the multifaceted roles of mitochondria, any impairment in mitochondrial function could be a mediator in the development or progression of cardiac disease. Cardiovascular diseases are the leading cause of mortality worldwide. The majority of cardiac pathologies such as myocardial infarction, heart failure, and ischemia/reperfusion (I/R) injury are usually attributed to the detrimental effects of an excessive increase in mitochondrial calcium and ROS generation [4]. Failure of mitochondrial calcium homeostasis leads to the opening of the mitochondrial permeability transition pore (mPTP), followed by the loss of ΔΨm, metabolic collapse, matrix swelling, and finally outer mitochondrial membrane (OMM) rupture, due to the enhanced permeability of the inner mitochondrial membrane (IMM) [5– 7]. Disruption of the OMM triggers apoptotic cascade by release of cytochrome c and apoptotic inducible factors, resulting in caspase-dependent and independent, respectively, irreversible cell damage and subsequently cardiac dysfunction [8]. Recent advances in our understanding of the role of mitochondria in the etiology and progression of cardiovascular diseases are based on multiple approaches to assess mitochondrial function and dysfunction. Isolation of mitochondria from animal issue is a powerful tool to investigate mitochondrial dysfunction in a disease condition. In this chapter, we aim to provide a detailed protocol for isolation of mitochondria followed by calcium retention capacity (CRC) measurement to assess mitochondrial Ca2+ handling. The first part of the chapter discusses step-by-step protocols involving isolation and purification of intact functioning mitochondria from guinea pig heart tissue using differential centrifugation. The same protocols can be adopted without modification to isolate mitochondria from rat cardiac tissue, as well. Furthermore, we describe the mitochondrial oxygen consumption rate (OCR) measurement using a Clark-type electrode to ascertain functional viability of the isolated mitochondria. In the second part of the chapter, we outline here a fluorescence based real-time measurement of CRC and mitochondrial membrane potential (ΔΨm) using the fluorometer and fluorescence Ca2+-sensing extra-mitochondrial dye Fura-4FF, and Tetramethylrhodamine, Methyl Ester, Perchlorate (TMRM), respectively.

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Materials 1. Heart Tissues: Albino guinea pigs (weight 250–400 g); use fresh guinea pig cardiac tissue. 2. Equipment and Supplies.

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(a) CAT X120 Homogenizer. (b) High speed refrigerated centrifuge (Eppendorf 5810 R High Speed Centrifuge) and 5804/R; 5810/R fixedangle rotor (Eppendorf FA-45-6-30), (c) Clark O2 electrode system (System S 200A; Strathkelvin Instruments, Glasgow, UK). (d) Fluorescence spectrophotometer (Qm-8, Photon Technology International, Horiba, Birmingham, NJ, USA). 3. Buffers: All solutions are freshly made. (a) Mitochondria Isolation buffer (MIB): 200 mM Mannitol, 50 mM Sucrose, 5 mM KH2PO4, 5 mM 3-(N-morpholino) propanesulfonic acid (MOPS), 1 mM Ethylene glycol-bis(β-aminoethyl ether)-N,N,N0 ,N0 -tetraacetic acid (EGTA), 0.1% Bovine serum albumin (BSA), pH 7.15 (adjusted with KOH). (b) Experimental buffer: 130 mM KCl, 5 mM K2HPO4, 20 mM MOPS, 0.1% BSA, pH 7.15 (adjusted with KOH). (c) 1 M sodium pyruvate, pH 7.4. (d) 2 mM Calcium chloride stock solution prepared in water. 4. Fluorophores. (a) Ratiometric calcium probes: Fura-4F pentapotassium salt (Kd 770 nM; λex: 340 and 380 nm, λem: 510 nm; Invitrogen™, Eugene, OR). Make 1 mM stock by dissolving in DMSO. Make aliquots and store at 20  C. (b) Mitochondrial membrane potential probe TMRM (λex: 546 and 573 nm, λem: 590 nm; Invitrogen™, Eugene, OR). Make 1 mM stock by dissolving in DMSO. Make aliquots and store at 20  C.

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3.1 Isolation of Mitochondria from Guinea Pig Heart Using Differential Centrifugation

This protocol is used for isolating mitochondria from laboratory animal, guinea pig, and rat [7, 9]. The key steps include: (1) tissue extraction, (2) homogenization, and (3) differential centrifugation (see Note 1). Animal handling is done according to procedures and guidelines of the National Institutes of Health, and the protocols were approved by the Institutional Animal Care and Use Committee of the Medical College of Wisconsin, Milwaukee, Wisconsin. Tissue Extraction

1. Euthanize the animal according to institutionally approved procedure. In this protocol, the guinea pig is anesthetized by intraperitoneal injection with 75 mg/kg ketamine along with the

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blood anticoagulant heparin 700 units. The guinea pig is euthanized by exsanguination via decapitation, when unresponsive to noxious stimulation, as per our institutionally approved guidelines of euthanasia (see Note 2). 2. Place the animal in the supine position on an animal surgical tray. Using surgical scissors and forceps, make an incision from the mid-abdomen to the diaphragm. Cut the sternum to open the chest cavity and expose the heart. Remove pericardium and excise the heart (see Note 3). 3. Transfer the excised heart to a 10 mL beaker containing ice cold MIB. Make small incisions and gently squeeze the heart to remove blood. Clean and rinse the explanted heart tissue for two more times to remove any residual/clotted blood. Trim any excess non-cardiac tissue, fat, and connective tissues using fine scissors (see Note 4). 4. Submerge the heart in small amount of buffer (~1 mL) and mince it into small pieces. Tissue Homogenization

5. Transfer the minced tissue with the isolation buffer into the 50 mL falcon tube, add 4 mL buffer. Quickly macerate the tissue using a homogenizer on the lowest speed for 10–15 s to break up any large chunks. 6. Add 2.65 mL of MIB + protease (5 U/mL of protease (Bacillus licheniformis)) and further homogenize for 10 s. Bring volume to 25 mL with MIB and homogenize for another 15–20 s to obtain uniform pale colored homogenate (see Note 5). Differential Centrifugation

7. Centrifuge the homogenate at 8000  g in 4  C for 10 min to remove the protease. 8. Resuspend the pellet initially in 12.5 mL of MIB and then further add 12.5 mL of the MIB (see Note 6). 9. Centrifuge the mitochondrial suspension at 800  g in 4  C for 10 min. 10. Carefully transfer the supernatant to a fresh 50 mL falcon tube, without disturbing the pellet. 11. Centrifuge the supernatant at 8000  g in 4  C for 10 min to obtain crude mitochondria pellet. 12. Discard the supernatant and resuspend the final mitochondrial pellet gently in a 200 μL of ice cold MIB (see Note 7). 13. Quantify the concentration of crude mitochondria by the Bradford method. Usually one guinea pig heart yields, 3–4 mg of mitochondria (see Note 8).

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Fig. 1 The measurement of functional integrity of the isolated mitochondria from a guinea pig heart. The time course of the rate of mitochondrial oxygen consumption measured using Clark type electrode during states 2, 3, and 4 respiration with complex I substrate, Pyruvate. The respiratory control index (RCI) was derived from the ratio of the rate of oxygen consumption/min for state 3 (in the presence of ADP) to state 4 (after exhausting most of the added ADP) 3.2 Measurement of Mitochondrial Respiration To Establish the Viability of the Isolated Mitochondria

The major challenge during the mitochondria isolation procedure is to preserve mitochondria structural integrity, which is imperatively associated with function. Mitochondrial respiration measurement is a well-established index of “healthy” mitochondrial populations. Here we describe the method involving polarographic measurement of oxygen consumption in the presence of substrates and ADP (Fig. 1). The protocol begins with measurement of state 1 respiration, when isolated mitochondria are resuspended in respiration medium containing oxygen and inorganic phosphate and consumes oxygen by oxidizing endogenous substrate. The addition of substrate depletes endogenous ADP and consumes residual oxygen (state 2 respiration). Furthermore, the addition of ADP in the presence of excess substrate maximally stimulates respiration with rapid depletion of oxygen (state 3 respiration). When all ADP is phosphorylated into ATP, the respiration rate reinstates to the pre-ADP addition values (state 4 respiration). The residual oxygen consumption after ADP depletion during state 4 respiration is largely due to H+ leak through the IMM [10]. The respiratory control index (RCI) ratio of states 3 to 4 respiration rates signifies the efficiency of mitochondria to utilize oxygen and nutrients to generate energy. Higher RCI suggests functional mitochondria, with the tight coupling of oxygen consumption to phosphorylation, while lower RCI indicates leaky IMM. 1. Calibrate the Clarke-type oxygen electrode. With our system the oxygen electrode was calibrated with air-saturated water (PO2 ~ 150 mmHg) and sodium sulfite (Na2SO3) solution (Sigma, St. Louis, MO; to achieve near zero PO2) (see Note 9).

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2. Add 250 μL of fresh experimental buffer and 0.25 mg mitochondria/mL. Start the recording of the oxygen consumption and wait for 1 min to obtain a stable baseline (state 1) (see Note 10). 3. Using 10 μL Hamilton microsyringe, add 5 μL of 1 M pyruvate to obtain a final concentration of 10 mM. This allows the determination of basal complex I-supported respiration (State 2) (see Note 11). 4. Following 1 min of stable oxygen consumption, add 5 μL of 250 mM ADP for a final concentration of 250 μM ADP to obtain maximal (state 3) mitochondrial respiration through complex I. 5. Wait for 6–8 min, the ATP-driven high oxygen consumption rate will transition back to baseline (state 4) once all the exogenously added ADP is phosphorylated to ATP. 6. The respiratory control index (RCI) is calculated as the state 3-to-state 4 ratio. For well-coupled mitochondria, the state 3 rate of respiration should be at least threefold greater than the state 4 rate of respiration (see Note 12). 3.2.1 Measurement of Calcium Retention Capacity of Mitochondria

Here, we describe methodology to measure the mitochondrial Calcium (Ca2+) retention capacity in isolated mitochondria, using a cuvette-based fluorescence spectrophotometer (Photon Technology International Inc.) (Fig. 2a). The level of Ca2+ outside mitochondria is assessed in these experiments, using Ca2+-sensing fluorophore in the extra-mitochondrial buffer. CaCl2 pulses are added to mitochondrial suspension until mPTP opening is observed. In response to each CaCl2 pulse, an increase in dye fluorescence intensity is observed, which then returns to a baseline as mitochondria take up and sequester the added calcium. When the concentration of free Ca2+ exceeds the Ca2+ buffering capacity of mitochondria, the mPTP opens which is marked by a dramatic rise in the fluorescence intensity in the extra-matrix buffer [7, 9]. 1. Resuspend 0.5 mg of isolated mitochondria in 1 mL of experimental buffer in a spectrofluometric cuvette under continuous stirring at room temperature (~23  C) (see Notes 13 and 14). 2. Add 1 μL of 1 mM Fura-4F pentapotassium salt stock, such that the final concentration is 1 μM (see Notes 15 and 16). 3. Start fluorescence measurements in a multi-wavelength-excitation dual wavelength-emission fluorimeter (Delta RAM, PTI) at 340- and 380-nm excitation and 535-nm emission for fura-4FF. 4. Add 10 mM pyruvate to the mitochondrial suspension at 30 s to energize mitochondria.

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Fig. 2 The measurement of Ca2+ retention capacity in the isolated mitochondria from a guinea pig heart. The time course of extra-mitochondrial Ca2+ measured with the fluorescent Ca2+ indicator probe Fura-4F (a) and change in ΔΨm (b) using fluorescence spectrophotometry. Mitochondria were energized with complex I substrate, Pyruvate at 60 s and 20 μM CaCl2 pulses were added every 300 s thereafter, in the absence (control, black trace) or in the presence of mPTP inhibitor, 0.5 μM Cyclosporin A (red tracing)

5. Add 10 μL aliquots of a 2 mM CaCl2 solution (final concentration 20 μM) every 5 min (300 s) until mPTP opening is observed (see Note 17). 6. The CRC is determined by converting the pulses to total amount of CaCl2 added (nmol) to the cuvette per mg of mitochondrial protein in the cuvette. Addition of 20 μM CaCl2 pulse (20 nmol/mL/0.5 mg/mL) results in sequestration of 40 nmol CaCl2 /mg mitochondrial protein. 3.2.2 Measurement of Calcium Induced ΔΨ m Loss in Isolated Mitochondria

A major driving force for mitochondrial Ca2+ uptake is a high potential gradient (ΔΨm) generated by proton translocation into the intermembrane space (IMS) by the respiratory electron chain complexes during oxidative phosphorylation. The sustained increase in mitochondrial free calcium either due to increased Ca2+ uptake or decreased efflux or failed sequestration can induce opening of the mPTP, leading to dissipation of ΔΨm [7, 9]. We describe here a dual excitation ratiometric approach using the fluorescent dye, TMRM to assess ΔΨm (Fig. 2b) (see Note 18). 1. Resuspend 0.5 mg of isolated mitochondria in 1 mL of experimental buffer in a spectrofluometric cuvette under continuous stirring at room temperature (~23  C) (see Notes 13 and 14). 2. For measurements of ΔΨm, add 1 μL of 1 mM TMRM in the mitochondrial suspension, such that the final concentration is 1 μM (see Note 16). 3. Start fluorescence measurements at 546 and 573 nm excitations, and 590 nm emission. 4. Add 10 mM pyruvate to the mitochondrial suspension at 30 s to energize mitochondria.

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5. Add 10 μL aliquots of a 2 mM CaCl2 solution (final concentration 20 μM) every 5 min (300 s) to introduce 200 nmol Ca2+/ mg mitochondrial protein until maximum membrane potential depolarization is reached (see Note 17). 6. The total CaCl2 added before membrane potential collapses is determined by converting the CaCl2 pulses to total amount of CaCl2 (nmol) added to the cuvette per mg of mitochondrial protein in the cuvette. Addition of 20 μM CaCl2 pulse (20 nmol/mL/0.5 mg/mL) results in addition of 40 nmol CaCl2/mg mitochondrial protein.

4

Notes 1. The entire procedure should be performed on ice, using pre-cool glassware’s and refrigerated equipment maintained at 4  C. 2. The appropriate guidelines set by the institution, in accordance with national regulations should be strictly followed, while handling animals. 3. The heart should be extracted and placed in cold MIB ideally within 15–20 s after decapitation. A quick extraction of the heart is critical to prevent degradation of mitochondria. 4. This step needs to be done as quickly as possible to prevent degradation of mitochondria. 5. Homogenize evenly at medium speed, but do not over homogenize the tissue as it may damage mitochondria. 6. To resuspend the pellet, use a disposable transfer pipette with a cut-out end of the tip. Make sure to completely break up all tissue chunks and mix well until the pellet is completely in suspension. 7. The volume of the resuspension depends on the pellet size and should be as small as possible. Resuspend the pellet by slowly pipetting up and down first with a cut 200 μL pipette tip to minimize shearing forces Avoid the formation of air bubbles during the resuspension process. 8. Other protein quantification methods can also be used (e.g., BCA). It is recommended to test several dilution factors (25, 50, and 100) to obtain an accurate sample concentration. 9. Calibration procedures vary from instrument to instrument. One should follow the manufacturer’s instructions for the instrument to be used. 10. Avoid introducing air bubbles into the chamber, as bubbles will displace volume, reducing the effective volume of the chamber.

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11. Usually, pyruvate is enough to stimulate complex I-specific respiration in guinea pig heart isolated mitochondria; for rat cardiac mitochondria, malate + pyruvate seem to be the preferred substrate. 12. The ratio of state 3 respiration rate to state 4 respiration rate is expressed as the respiratory control index (RCI) and should be greater than 7.0 for complex I-specific respiration for wellcoupled wild-type mitochondria. An RCI of less than 5.0 for mitochondria most commonly suggests a suboptimal concentration of mitochondria or a technical problem with the isolation of mitochondria that damages the integrity of the inner membrane. 13. Isolated mitochondria kept on ice show preserved bioenergetics (tight RCI and ΔΨm) and CRC for 8–9 h. 14. Though the experimental buffer contains no EGTA, since the mitochondria are suspended in isolation buffer that contains 1 mM EGTA, a residue of EGTA is carried over with aliquot of mitochondria transferred to experimental buffer resulting in an approximately 40 μM EGTA in 1 mL of mitochondria suspension. 15. This salt form of the fluorescent dye is impermeable to mitochondria and measures the amount of calcium outside mitochondria. 16. Protect the experimental buffer from light following the addition of the dye. 17. The amount and timing of the CaCl2 injections are defined by the experimenter and can be adjusted for according to experimental needs. 18. Measurement of calcium induced loss in ΔΨm is an alternative approach to measuring the CRC, however, this method does not provide the additional information on the kinetics of Ca2+ fluxes.

Acknowledgments We would like to acknowledge James Heisner for setting up and optimizing the approaches described here. This work was supported by a grant from NIH: R01 HL131673 to AKSC. References 1. Page E, McCallister LP (1973) Quantitative electron microscopic description of heart muscle cells. Application to normal, hypertrophied

and thyroxin-stimulated hearts. Am J Cardiol 31:172–181

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2. Harris DA, Das AM (1991) Control of mitochondrial ATP synthesis in the heart. Biochem J 280(Pt 3):561–573 3. Jouaville LS, Pinton P, Bastianutto C, Rutter GA, Rizzuto R (1999) Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic priming. Proc Natl Acad Sci U S A 96:13807–13812 4. Camara AK, Bienengraeber M, Stowe DF (2011) Mitochondrial approaches to protect against cardiac ischemia and reperfusion injury. Front Physiol 2:13 5. Bernardi P (1999) Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79:1127–1155 6. Camara AK, Lesnefsky EJ, Stowe DF (2010) Potential therapeutic benefits of strategies directed to mitochondria. Antioxid Redox Signal 13:279–347

7. Mishra J, Davani AJ, Natarajan GK, Kwok WM, Stowe DF, Camara AKS (2019) Cyclosporin a increases mitochondrial buffering of calcium: an additional mechanism in delaying mitochondrial permeability transition pore opening. Cell 8:1052 8. Crompton M (1999) The mitochondrial permeability transition pore and its role in cell death. Biochem J 341(Pt 2):233–249 9. Natarajan GK, Glait L, Mishra J, Stowe DF, Camara AKS, Kwok WM (2020) Total matrix ca(2+) modulates ca(2+) efflux via the Ca(2+)/ H(+) exchanger in cardiac mitochondria. Front Physiol 11:510600 10. Agarwal B, Dash RK, Stowe DF, Bosnjak ZJ, Camara AK (2014) Isoflurane modulates cardiac mitochondrial bioenergetics by selectively attenuating respiratory complexes. Biochim Biophys Acta 1837:354–365

Chapter 7 Characterizing the Electron Transport Chain: Structural Approach Ting Liang, Janice Deng, Bijaya Nayak, Xin Zou, Yuji Ikeno, and Yidong Bai Abstract The mitochondrial respiratory chain which carries out the oxidative phosphorylation (OXPHOS) consists of five multi-subunit protein complexes. Emerging evidences suggest that the supercomplexes which further consist of multiple respiratory complexes play important role in regulating OXPHOS function. Dysfunction of the respiratory chain and its regulation has been implicated in various human diseases including neurodegenerative diseases and muscular disorders. Many mouse models have been established which exhibit mitochondrial defects in brain and muscles. Protocols presented here aim to help to analyze the structures of mitochondrial respiratory chain which include the preparation of the tissue samples, isolation of mitochondrial membrane proteins, and analysis of their respiratory complexes by Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) in particular. Key words Blue Native Gel, Respiratory complex, Assembly, Brain, Muscle

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Introduction The primary function of mitochondria is to generate ATP through oxidative phosphorylation (OXPHOS) process. The OXPHOS happens along the respiratory chain resided at the inner membrane of mitochondria [1]. Emerging evidence supports a model proposing that the MRC functions not via isolated protein complexes floating in the inner mitochondrial membrane, but rather as highly regulated multicomplex structures [2, 3]. Mutations in the subunits/assembly factors of the respiratory machinery have been associated with many human diseases [4, 5]. More importantly, mitochondrial dysfunction including defects in respiratory chain complexes have been recorded in expanding numbers of human diseases including neurodegenerative diseases, metabolic diseases, and cancer [6–9]. Accordingly, numerous mouse models have been developed which exhibits mitochondrial defects [10–12]. As a

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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result, demand for more reliable methods of assessing mitochondrial respiratory chain, in particular for determination of the dynamics of respiratory complexes in tissue samples has significantly increased. Mitochondrial inner membrane is enriched with protein complexes involved in multiple biological processes including OXPHOS [1]. Blue Native (BN)-PAGE, in contrast to sodium dodecyl sulfate (SDS)-PAGE, was designed to resolve protein complexes in native conditions and to preserve the protein–protein interactions. With a proper gradient concentration, it is particularly useful for determination of membrane protein complexes in the mass range of 10 kDa to 10 MDa [13]. Mitochondrial respiratory complexes just fall in this range. The improvement of extraction of respiratory complexes from the mitochondrial inner membrane is facilitated by identifications of nonionic detergents which are used for the solubilization of biological membranes for different purposes. Digitonin, the mildest detergents, has been used to isolate respiratory supercomplexes consist of multiples respiratory complexes. Dodecylmaltoside (DDM), with a stronger delipidating property, is used for isolation of individual complexes. Triton X-100 sometimes is used to show intermediate behaviors when a complementary approach is desired [14–16]. Here we only utilize digitonin as the detergent as we are particularly interested in the characterization of respiratory supercomplexes. Quantitative and qualitative analysis of respiratory complexes can be followed with in-gel activity staining procedures for the enzymatic activities associated with a particular respiratory complex; and immunoblotting on 1D BN-PAGE for representative subunits for each of the respiratory complexes. The determination of the distribution of individual complexes, supercomplexes which contain multiple respiratory complexes, and high molecular weight megacomplexes containing two complex I together with other complexes would probably shed light on the regulation of energy metabolism and redox signaling.

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Materials

2.1 Isolation of Mitochondrial Complex (See Note 1)

1. Mitochondrial solubilization buffer (SB): 50 mM NaCl, 50 mM BisTris/HCl pH 7.0, 2 mM 6-aminohexanoic acid (6-ACA), 1 mM EDTA, adjust pH by HCl to 7.4, store at 4  C. 2. 5% Bovine Serum Albumin Fatty Acid-Free (BSA-FF): Weigh 5 g BSA-FF and dissolve in SB to a final volume of 100 ml, store at 20  C. 3. 20% Digitonin: Weigh 0.2 g digitonin and make to 1 ml distilled water, heat at 95  C–98  C to dissolve, keep at room

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temperature to cool, and then centrifuge at 20,000  g for 1 min at 4  C, retain the supernatant, store at 20  C. 4. Protease Inhibitor (100 X PI): Reconstitute each vial with 1 ml H2O to obtain a 100 stock solution. 5. 0.25% Trypsin: dissolve 10 ml 2.5%Trypsin into 90 ml distilled water, store at 20  C. 6. Buffer M: Mix 19.1 g mannitol and 11.9 g sucrose, 5 ml 1 M Tris/HCl pH 7.4, 1 ml 0.5 M EDTA pH 8.0 into distilled water to a final volume of 500 ml, filter, Store at 4  C (see Note 2). 7. Buffer M1: Add 5 ml 5% BSA-FF, 50 μl 20% digitonin, 500 μl PI 100 into Buffer M to a final volume of 50 ml, prepare fresh, Store in 20  C, good for 1 week. 8. Buffer M2: Add 10 ml 0.25%Trypsin, 20 μl 0.5 M EDTA pH 8.0 into Buffer M1 to a final volume of 50 ml, prepare fresh, Store at 20  C, good for 1 week. 2.2 Gradient Gel Electrophoresis

1. 4 Gel Buffer: 0.2 M 6-ACA, 150 mM BisTris/HCI pH 7.0, store at 4  C. 2. 50% Glycerol: Mix 25 ml 100% Glycerol into 25 ml distilled water, store at 4  C. 3. 30% Acrylamide/Bis Solution, 37.5:1. 4. 10% Ammonium persulfate (APS), prepare fresh each day. 5. TEMED, keep in the dark at 4  C. 6. Isopropanol. 7. 3% separation gel: Mix 6.5 ml water, 2.5 ml 4  gel buffer, 1 ml 30% Acrylamide/Bis Solution, 50 μl 10% APS, and 5 μl TEMED; 10 ml per gel. 8. 11% separation gel: Mix 0.88 ml 4  gel buffer, 1.3 ml 30% Acrylamide/Bis Solution, 1.3 ml 50% Glycerol, 15 μl 10% APS, and 1.5 μl TEMED; 3.5 ml per gel. 9. 3% stacking gel: Mix 1.9 ml water, 0.75 ml 4  gel buffer, 0.33 ml 30% Acrylamide/Bis Solution, 30 μl 10% APS, and 5 μl TEMED; 3 ml per gel. 10. Comb with 1.5 mm wide teeth, 10 wells. 11. Peristaltic pump. 12. 5% Coomassie Brilliant Blue G-250: Add 0.5 g Coomassie Brilliant Blue G-250, 0.6559 g 6-ACA into distilled water to a final volume of 10 ml, store at 4  C. 13. 80% Glycerol: Mix 40 ml 100% Glycerol into10 ml distilled water, store at 4  C. 14. Loading buffer: Mix 5 ml 5% Coomassie Brilliant Blue G-250 into 5 ml 80% Glycerol, store at 4  C.

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15. Cathode Buffer: Mix 8.96 g Tricine, 0.2 g Coomassie Brilliant Blue G-250, 15 ml 1 M BisTris/HCl pH 7.0 into distilled water to a final volume of 1 L, store at 4  C. 16. Colorless Cathode Buffer: Mix 8.96 g Tricine, 0.02 g Coomassie Brilliant Blue G-250, 15 ml 1 M BisTris/HCl pH 7.0 into distilled water to a final volume of 1 L, store at 4  C. 17. Anode Buffer: Mix 50 ml 1 M BisTris/HCl pH 7.0 into distilled water to a final volume of 1 L, store at 4  C. 2.3

Immunoblotting

1. PVDF membrane. 2. 3 mm filter paper. 3. Transfer buffer: Mix 3 g Tris Base, 14.4 g Glycine, 100 ml methanol into distilled water to a final volume of 1 L, store at 4  C. 4. Wash buffer (TBST): Mix 20 ml 1 M BisTris/HCl pH 7.0, 30 ml 5 M NaCl, 1 ml Tween-20 into 1 L distilled water. 5. Blocking solution: 5% milk in TBST. 6. Primary antibody: Grim19 (for complex I), COXIV (for complex IV). 7. Secondary antibody: AP-conjugated secondary antibodies. 8. Stock developing buffer: Mix 6.05 g Tris Base, 0.25 ml 1 M MgCl, 200 μl HCl into distilled water to a final volume of 500 ml, store at 4  C. 9. 50 mg/ml Nitro blue tetrazolium (NBT): Dissolve 0.05 g NBT in 70% dimethylformamide (DMF) to 1 ml, store at 20  C. 10. 50 mg/ml 5-bromo-4-chloro-3- indolyl phosphate (BCIP): Dissolve 0.05 g BCIP in 100% DMF to 1 ml, store at 20  C. 11. Developing solution: Mix 52.8 μl NBT, 26.4 μl BCIP into 8 ml stock developing buffer, prepare fresh; 8 ml per membrane.

2.4

Gel Assay

1. Complex I staining buffer: Mix 0.01 g NBT, 0.001 g NADH, 1 ml 1 M Tris/HCl pH 7.4 into distilled water to a final volume of 10 ml, prepare fresh. 2. Complex IV staining buffer: Mix 0.005 mg Diaminobenzidine tetrahydrochloride (DAB), 0.75 g Sucrose, 0.01 g Cytochrome c, 0.38 ml 0.5 M NaH2PO4, 1.62 ml 0.5 M Na2HPO4 into distilled water to a final volume of 10 ml, prepare fresh.

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Methods

3.1 Isolation of Mitochondrial from Mice Tissues (See Note 3)

1. Whole brain is taken out of cranial cavity and then placed on Precision Brain Slicer, on ice. The brain is subsequently cut into half at the center of brain (Fig. 1a). The left half of the brain is utilized to isolate mitochondria. Skeletal muscles are taken from the hind legs (Fig. 1b). Each muscle is cut into small pieces on a 10-mm dish on ice, and then are rinsed several times with PBS. 2. Transfer tissues to 2 ml homogenizer in 2 ml buffer M1 or buffer M2 (see Note 4). Homogenize tissue 10–15 strokes by pestle and keep in 1.5 ml tubes (see Note 5). Sit on ice for 10 min. 3. The homogenate is then centrifuged at 20,000  g for 10 min to obtain a pellet containing nuclei, mitochondria, and larger cell fragments. Remove supernatant totally. Pellets can be stored at- 80  C or continue with the following step. 4. The pellet is weighed in balance and obtain the wet weight tissues, the pellet is then resuspended with 35 μl SB to 15 mg weight skeletal muscle, 10 mg weight brain (see Note 6). 5. Then add 10 μl 20% digitonin for skeletal muscles, 20 μl 20% digitonin for brain. 6. Solubilize for 5–10 min on ice. 7. Centrifuge at 22,000  g for 60 min at 4  C. Retain the supernatant. 8. Determine the concentration of protein using the Pierce BCA protein assay kit.

3.2 Gel Electrophoresis

1. Cast 3–11% gradient separation gels using Peristaltic pump. 2. Lay 0.6 ml isopropanol over the gel to straighten the surface. 3. Polymerization takes approximately 40–50 min. 4. Discard the isopropanol and wash in distilled water to remove residual isopropanol. 5. Load 3% stacking gel, and insert the 10 wells comb. 6. Polymerization takes approximately 30 min, store the gels at 4  C or follow the next step. 7. Before loading the samples, fill the wells with the blue cathode buffer and wash any excess non-polymerized acrylamide from the well. 8. Prepare 30 μg protein to detect supercomplex I, 40 μg protein to detect supercomplex IV. Mix 1/5 volumes of protein loading buffer to the samples, load in wells for BN-PAGE.

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Fig. 1 Collect and process of brain and muscle samples. (a) Brain. (b) Muscle

9. Fill the gel cathode chamber with the blue cathode buffer and the bottom chamber with the anode buffer. 10. Run the gel at constant Ampere of 5 mA/gel for about 1 h or until the dye front has traveled approximately 1/3 of the gel. Then, change the cathode buffer to colorless cathode buffer and run at same Ampere for about 2–3 h until the most of blue dye leaves the gel (see Notes 7 and 8). 3.3

Immunoblotting

1. Wash the gel in transfer buffer to remove dye (see Note 9). 2. Activate the PVDF membrane in methanol for 1 min and then immerse into the transfer buffer. Pre-wet the filter paper and sponge in transfer buffer (see Note 10). 3. Assemble the transfer sandwich (sponge, filter paper, membrane, gel, filter paper, sponge) and then put the cassette, gel transfer is performed at 90 V about 90 min at 4  C. 4. Following transfer, rinse PVDF membrane in TBST for 10 min, 2–3 times. 5. Block the membrane with 5% milk for 1 h. 6. Rinse the membrane in TBST and incubate with primary antibody (Grim19, COXIV) for 2 h at room temperature or overnight at 4  C. All antibodies are diluted to 1:1000 in TBST. 7. Wash the membrane in TBST for 10 min, 3–4 times. 8. Incubate the membrane with second antibody for 1 h at room temperature. Antibody is diluted to 1:5000 in TBST. 9. Wash the membrane in TBST for 10 min, 3–4 times. 10. Immerse the membrane into developing solution until the bands are observed, then transfer it to the water, quick immersion in 100% methanol until most of dye has been removed and put it to the water, keep in the TBST. 11. Scan the stained membrane, as shown in Fig. 2.

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Fig. 2 Analysis of mitochondrial respiratory complexes with immunoblotting. (a) Western Blot of CI (Grim 19) after BN-PAGE with the mice tissues of brain(B) and muscle(M). (b) Western Blot of CIV (COXIV) after BN-PAGE with the mice tissues of brain (B) and muscle(M). Megacomplexes (MCs), supercomplexes (SCs)

Fig. 3 Analysis of mitochondrial respiratory complexes with Gel Assay. (a) CI (Grim 19) in gel activity assay after BN-PAGE with the mice tissues of brain(B) and muscle(M). (b) CIV (COX IV) in gel activity assay after BN-PAGE with the mice tissues of brain(B) and muscle(M). Megacomplexes (MCs), supercomplexes (SCs)

3.4

Gel Assay

1. After electrophoresis separation by the method described above (see Subheading 3.2), the respiratory complexes retain enzymatic activity. 2. Incubate the gel into complex I staining buffer for 30–40 min at room temperature, the blue-purple color can develop as shown in Fig. 3a. 3. Incubate the gel into complex IV staining buffer about 2–3 h at room temperature, brown color can develop as shown in Fig. 3b. 4. Wash the gel completely in water (see Note 11). 5. Scan the stained gels.

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Notes 1. All solutions used should be sterile and nuclease-free. 2. Heating promotes mannitol dissolve, measure pH after it cools down to room temperature. 3. All steps for insolation of mitochondria are performed on ice. 4. Buffer M1 is used for brain, liver and kidney samples, Buffer M2 is used for heart and muscle samples. 5. Pre-cool tubes, homogenizer, and buffer M1/M2 on ice, preset the centrifuge at 4  C. 6. Gently and carefully resuspend the pellet to reduce exposure to oxygen. 7. Excess Coomassie blue dye will interfere with western blots and gel activity assays, so change to colorless cathode buffer in time. 8. Run gels at 4  C to avoid overheating. 9. Cut the blue dye completely if it still keeps in the gel, Coomassie will interfere with the antibody binding thus resulting in high background. 10. Make sure the sponge and filter paper are evenly and extensively saturated, Air bubbles in the sponge or filter paper result in areas where the protein does not transfer well to the membrane. 11. Putting Kimwipes in the water will help to absorb residual reagents present in the gel and will eliminate background on gels faster.

Acknowledgments This work is supported by grants from National Institute of Health (R01 GM109434 and GM130129), the Pathology Core in the San Antonio Nathan Shock Center (P30-AG013319), and LT and YB are also supported by William and Ella Owens Medical Research Foundation. References 1. Saraste M (1999) Oxidative phosphorylation at the fin de sie`cle. Science (New York, NY) 283(5407):1488–1493. https://doi.org/10. 1126/science.283.5407.1488 2. Porras CA, Bai Y (2015) Respiratory supercomplexes: plasticity and implications. Front Biosci (Landmark Ed) 20:621

3. Vartak R, Porras CA-M, Bai Y (2013) Respiratory supercomplexes: structure, function and assembly. Protein Cell 4(8):582–590 4. Guerrero-Castillo S, Baertling F, Kownatzki D, Wessels HJ, Arnold S, Brandt U, Nijtmans L (2017) The assembly pathway of mitochondrial respiratory chain complex I. Cell Metab 25(1):128–139

Respiratory Complexes Assembly Determination in Tissue Samples 5. Barrientos A, Gouget K, Horn D, Soto IC, Fontanesi F (2009) Suppression mechanisms of COX assembly defects in yeast and human: insights into the COX assembly process. Biochim Biophys Acta 1793(1):97–107. https:// doi.org/10.1016/j.bbamcr.2008.05.003 6. Bhatti JS, Bhatti GK, Reddy PH (2017) Mitochondrial dysfunction and oxidative stress in metabolic disorders—A step towards mitochondria based therapeutic strategies. Biochim Biophys Acta Mol Basis Dis 1863(5): 1066–1077 7. Trushina E, McMurray CJN (2007) Oxidative stress and mitochondrial dysfunction in neurodegenerative diseases. Neuroscience 145(4): 1233–1248 8. Schapira A, Gu M, Taanman JW, Tabrizi S, Seaton T, Cleeter M, Cooper JM (1998) Mitochondria in the etiology and pathogenesis of Parkinson’s disease. Ann Neurol 44(S1 1): S89–S98 9. Haas RH (2019) Mitochondrial dysfunction in aging and diseases of aging. Multidisciplinary Digital Publishing Institute, Basel 10. Chesselet M-F, Richter FJTLN (2011) Modelling of Parkinson’s disease in mice. Lancet Neurol 10(12):1108–1118

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11. Eckert A, Schulz KL, Rhein V, Go¨tz J (2010) Convergence of amyloid-β and tau pathologies on mitochondria in vivo. Mol Neurobiol 41(2-3):107–114 12. Vempati UD, Torraco A, Moraes CTJM (2008) Mouse models of oxidative phosphorylation dysfunction and disease. Methods 46(4): 241–247 13. Wittig I, Braun H-P, Sch€agger H (2006) Blue native PAGE. Nat Protoc 1(1):418–428. https://doi.org/10.1038/nprot.2006.62 14. McKenzie M, Lazarou M, Thorburn DR, Ryan MT (2006) Mitochondrial respiratory chain supercomplexes are destabilized in Barth syndrome patients. J Mol Biol 361(3):462–469 15. Eubel H, J€ansch L, Braun H-P (2003) New insights into the respiratory chain of plant mitochondria. Supercomplexes and a unique composition of complex II. Plant Physiol 133(1):274–286. https://doi.org/10.1104/ pp.103.024620 16. Sch€agger H, Pfeiffer K (2000) Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J 19(8): 1777–1783. https://doi.org/10.1093/ emboj/19.8.1777

Chapter 8 Characterizing the Electron Transport Chain: Functional Approach Using Extracellular Flux Analyzer on Mouse Tissue Samples Ting Liang, Jay Dunn, Xin Zou, Bijaya Nayak, Yuji Ikeno, Lihong Fan, and Yidong Bai Abstract The Seahorse Extracellular Flux Analyzer enables the high-throughput characterization of oxidative phosphorylation capacity based on the electron transport chain organization and regulation with relatively small amount of material. This development over the traditional polarographic Clark-type electrode approaches make it possible to analyze the respiratory features of mitochondria isolated from tissue samples of particular animal models. Here we provide a description of an optimized approach to carry out multiwell measurement of O2 consumption, with the Agilent Seahorse XFe96 analyzer on mouse brain and muscles to determine the tissue-specific oxidative phosphorylation properties. Protocols include the preparation of the tissue samples, isolation of mitochondria, and analysis of their function; in particular, the preparation and optimization of the reagents and samples. Key words Seahorse, Electron transport chain, O2 consumption, Brain, Muscle

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Introduction Mitochondria have been more and more appreciated as functional regulators of life and death, yet still their primary function is to generate ATP through oxidative phosphorylation [1]. The mitochondrial respiratory chain (MRC) is composed of five protein complexes: NADH-ubiquinone oxidoreductase as Complex I, succinate-ubiquinone oxidoreductase as Complex II, ubiquinonecytochrome-c oxidoreductase as Complex III, cytochrome-c oxidase as Complex IV, and ATP synthase as complex V [2]. The mitochondrial respiratory chain (MRC) couples electron transfer with ATP production [3]. Emerging evidence supports a model proposing that the MRC functions not via isolated protein complexes floating in the inner mitochondrial membrane, but rather as highly regulated multicomplex structures, and such regulation

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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plays an important role in health and diseases [2, 4]. Thus, measurement of rates of O2 consumption are fundamentally informative, as electron transport and oxidative phosphorylation reflect the concerted mitochondrial function which further regulates various cellular processes. Mitochondrial dysfunction is implicated in the etiology of many common diseases including metabolic disorders, neurodegenerative diseases, and cancers, as well as the process of aging [5]. Defects in respiratory complexes, including those caused by mutations in the subunits/assembly factors of the respiratory machinery, have been observed in various clinical phenotypes [6– 9]. Recently various mouse models for such human diseases have been developed and characterized [10–13]. As a result, demand for more flexible and higher throughput methods of assessing mitochondrial function, in particular for determination of the electron transfer chain and oxidative phosphorylation properties in tissue samples has significantly increased. Although oxygen consumption measurements have been the preferred method of assessing mitochondrial function [14, 15], the low-throughput and low sensitivity nature of traditional polarographic Clark-type oxygraphy approaches have limited the applicability of such approach, particularly in animal tissue samples. Brain and muscle are among the most metabolically active tissues and consequently are most affected by mitochondrial dysfunction. Moreover, they also exhibit distinctive bioenergetics and physiological phenotypes [16]. An effective approach to analyze the respiratory features in these critical tissues are highly desired. The Agilent Seahorse XFe96 analyzer provided a possibility with high throughput with the sample volumes much lower than conventional polarographic analysis [17]. Isolation of mitochondria from tissues facilitates full access of substrates to the respiratory chain. As a result, direct interrogation of each component of the electron transfer chain is possible, as substrates and inhibitors can be added under careful control. To assess the tissue-specific organization and regulation of electron transfer chain, the respiration properties of the mitochondria isolated from brain and muscle as the representatives are investigated. In particular, Basal with ADP induced, Oligomycin inhibited, maximal respiration with FCCP induced, and Antimycin A inhibited oxygen consumption rate (OCR) have been sequentially measured in each well through the addition of specific reagents. Various regulatory properties of the respiratory chain including proton leakage, oxidative phosphorylation coupling, and the capacity of regulation on oxidative phosphorylation can be further determined.

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Materials

2.1 Surgical Instruments

1. Metzenbaum scissors. 2. Bone cutting straight scissors. 3. Narrow pattern serrated and curved forceps. 4. Precision Brain Slicer with sagittal sections.

2.2 Media and Reagents

1. 1 M Sucrose: Weigh 17.115 g sucrose and dissolve in distilled water to a final volume of 50 ml, filter, store at 4  C. 2. 1 M Mannitol: Weigh 9.11 g Mannitol and dissolve in distilled water to a final volume of 50 ml, filter, store at 4  C. 3. 1 M HEPES: Weigh 11.92 g HEPES and dissolve in distilled water to a final volume of 50 ml, adjust pH with KOH to 7.2, filter, store at 4  C. 4. 0.5 M EGTA: Weigh 9.51 g EGTA and dissolve in distilled water to a final volume of 50 ml, adjust pH with KOH to 8.0, filter, store at 4  C. 5. 5% Bovine Serum Albumin Fatty Acid-Free (BSA): Weigh 0.5 g BSA and dissolve in distilled water to a final volume of 10 ml, filter, store at 20  C. 6. 1 M KH2PO4: Weigh 6.8 g KH2PO4 and dissolve in distilled water to a final volume of 10 ml, filter, store at 4  C. 7. 1 M MgCL2: Weigh 4.76 g MgCL2 and dissolve in distilled water to a final volume of 10 ml, filter, store at 4  C. 8. 1 M ADP: Weigh 4.712 g ADP and dissolve in distilled water to a final volume of 10 ml, filter, store at 20  C. 9. 0.5 M Succinic acid: Weigh 0.59 g Succinic acid and dissolve in distilled water to a final volume of 10 ml, filter, store at 20  C. 10. 10 mM Oligomycin (stock): Weigh 7.91 mg Oligomycin and dissolve in 95% ethanol to a final volume of 1 ml, store at 20  C. 11. 31.6 μM Oligomycin (loading): 9.5 μl 10 mM Oligomycin with 2990.5 μl MAS without BSA to a final volume of 3 ml. Prepare fresh (see Note 1). 12. 10 mM FCCP (stock): Weigh 2.54 mg FCCP and dissolve in 95% ethanol to a final volume of 1 ml, store at 20  C. 13. 40 μM FCCP (loading): 12 μl 10 mM FCCP with 2988 μl MAS without BSA to a final volume of 3 ml. Prepare fresh (see Note 1). 14. 10 mM Antimycin A (stock): Weigh 5.49 mg Antimycin and dissolve in 95% ethanol to a final volume of 1 ml, store at 20  C.

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15. 40 μM Antimycin A (loading): 12 μl 10 mM Antimycin A with 2988 μl MAS without BSA to a final volume of 3 ml. Prepare fresh (see Note 1). 16. 10 mM Rotenone: Weigh 3.94 mg Rotenone and dissolve in 95% ethanol to a final volume of 1 ml, store at 20  C. 17. Mitochondrial isolation buffer: 70 mM Sucrose, 210 mM Mannitol, 5 mM HEPES, 1 mM EGTA, 0.2% BSA, adjust pH with KOH to 7.2, and store at 4  C (see Note 2). 18. Mitochondrial assay solution (MAS): 70 mM Sucrose, 220 mM Mannitol, 10 mM KH2PO4, 5 mM MgCL2, 2 mM HEPES, 1 mM EGTA, 0.2% BSA, adjust pH with KOH to 7.2, and store at 4  C (see Note 2). 19. Mitochondrial dilution/Complex II substrate buffer: 10 mM Succinate, 2 μM Rotenone, 4 mM ADP, 0.2% BSA, adjust pH with KOH to 7.2 (see Note 2). 2.3 Bioenergetics Measurement

1. Equipment: Agilent Seahorse XFe96 Extracellular Flux Analyzer. 2. Reagents: Seahorse XFe96 FluxPaks (sensor cartridges, cell culture plates, and calibrant buffer).

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Methods

3.1 Tissue Sample Preparation

1. The mice are euthanized with isoflurane in accordance with IACUC procedures. 2. Skeletal muscles are dissected from the hind legs. Each muscle is cut into small pieces on a 10-mm dish on ice, and rinse several times with PBS. 3. Whole brain is dissected out of cranial cavity and place on Precision Brain Slicer, which is placed on ice. The brain is cut into half at the center of brain. The right half of the brain is utilized to isolate mitochondria and measure respiration.

3.2 Mitochondria Isolation

1. The above tissue samples are homogenized in 10 volumes of mitochondrial isolation buffer using the tissue homogenizer for 8–10 strokes on ice, depending on the amount of tissue (see Notes 3 and 4). 2. Tissue homogenates are then centrifuged at 800  g for 8 min at 4  C. 3. Following the first centrifugation, the supernatant is collected and subjected to second centrifugation at 800  g for 8 min at 4  C.

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4. The mitochondrial fractions are then harvested by centrifugation of the above remaining supernatant at 8000  g for 10 min at 4  C. 5. The pellets are resuspended with mitochondrial isolation buffer and centrifuged at 15,000  g for 10 min at 4  C. 6. The final pellet is resuspended in a minimal volume of mitochondrial isolation buffer (see Note 5). 7. The protein concentration is then determined using the Pierce BCA protein assay kit. 3.3 Extracellular Flux Analysis

Day 0

1. 20 ml of Seahorse XF Calibrant in a conical tube are kept in a non-CO2 37  C incubator overnight. 2. The XFe96 Extracellular Flux Assay sensor cartridge package is opened, and 200 μl of sterile water per well is added to utility plate, and the sensor cartridge is placed back on the utility plate with the plate lid, and hydrated overnight in a non-CO2 37  C incubator. Day 1 Preparation of Respiration Reagents

3. The water is removed from the wells of the utility plate, and replaced with 200 μl per well of pre-warmed XF Calibrant, submerging the sensors. 4. The kit is kept in a non-CO2 37  C incubator for 45–60 min prior to loading the injection ports of the sensor cartridge. 5. After that respiration reagents are loaded as follows: port A, 20 μl of 31.6 μM Oligomycin (3.16 μM, final); port B, 22 μl of 40 μM FCCP (4 μM, final); port C, 24 μl of 40 μM Antimycin (4 μM, final), before inserting into the XFe96 (see Note 6). Mitochondrial Sample Preparation

6. Mitochondrial from brain and muscle are seeded in 25 μl of MAS into the wells of the Seahorse XFe96 cell culture Microplate. Background wells contain MAS only. 7. The microplate is then centrifuged in a swinging bucket microplate adaptor, at 2400  g for 20 min at 4  C. 8. After centrifugation, 155 μl of pre-warmed (at 37  C) mitochondrial dilution/substrate buffer is added to each well (see Notes 7 and 8). 9. The plate is then loaded to the Agilent Seahorse XFe96 Analyzer for initiation of the measurement.

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Notes 1. Loading of respiration reagent should be prepared fresh on the day of each experiment. 2. Adjust the pH of all solutions to 7.2 to maintain viable mitochondria. 3. Preset the centrifuge at 4  C, and pre-cool the mitochondrial isolation buffer and homogenizer, and tubes on ice. 4. The tissues are kept on ice throughout the isolation procedure. 5. Try to avoid introducing air bubbles when resuspending the mitochondria to minimize exposure to oxygen. 6. Gently and vertically load respiration reagents in ports to make sure the reagents are evenly filled for each well using a multichannel pipette. 7. Be careful when loading 25 μl mitochondria sample and 155 μl substrate buffer, avoiding air bubbles in wells. 8. Keep mitochondrial preparations on ice during processing which protects the integrity of the mitochondria, however, the preparations should be warmed up to 37  C before inserting the measurement plate into the Seahorse XFe96, to achieve the optimal condition for measurement. 9. In particular, for brain samples, as shown in Fig. 2, we find that the optimal amounts of brain mitochondria in this assay is between 2 and 5 μg of protein. Similarly, the same trend is recorded for increased OCR following FCCP injection (Fig. 3). In muscle samples, as shown in Fig. 4, we find that the optimal amounts of muscle mitochondria in this assay is between 0.3 and 3 μg. Similarly, the same trend is recorded for increased OCR following FCCP injection (Fig. 5). 10. As shown in Fig. 6, we find the maximal inductions with FCCP are achieved with 5 μM in brain preparations and 4 μM in muscle samples. Interestingly, the uncoupled rate is reduced when excessive amount of FCCP is used in this system.

5

Optimization of Assay Conditions in Mitochondria Isolated from Brain and Muscle Mitochondrial dysfunction has been implicated in various human diseases, most notably in brain and muscle, and various genetic and pharmacological mouse models with related mitochondrial defects mimicking such diseases have been established. It is, therefore, important to develop a reliable approach to follow the various features of mitochondrial respiratory capacity in brain and muscle with relatively limited amounts of materials. We previously

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Fig. 1 Determination of respiratory features with different amounts of brain and muscle samples. (a) 0.3–7 μg of brain samples are treated with ADP (4 mM), Oligomycin (3.16 μM), FCCP (4 μM), and Antimycin A (4 μM), respectively. (b) 0.2–7 μg of muscle samples are treated with ADP (4 mM), Oligomycin (3.16 μM), FCCP (4 μM), and Antimycin A (4 μM), respectively

demonstrated that brain and muscle exhibit tissue-specific regulation of respiration [16, 18]. We thus set out to establish the optimal conditions for investigating the regulation of OXPHOS in brain and muscle mitochondria in the Seahorse XFe96. Based on the results from a pilot experiments, we find the amounts of isolated mitochondria and concentration of FCCP are important to get an accurate and reproducible measurement. 5.1 Optimize the Amounts of Isolated Mitochondria (Figs. 1, 2, 3, 4, and 5)

1. Run assays of the brain samples with a range of 0.3–7 μg of mitochondrial protein in each plate, and muscle samples with a range of 0.2–7 μg of mitochondrial protein in each plate, respectively, to identify the optimal conditions for measurements. 2. The determinations of respiratory features including ADP induced, Oligomycin sensitive, FCCP induced, and Antimycin A sensitive OCR of different amounts of mitochondria from brain and muscle samples are shown in Figs. 1, 2, 3, 4, and 5 (see Note 9).

5.2 Optimize the Concentration of FCCP (Fig. 6)

We are puzzled by the fact that we could not achieve the maximal respiration induced by FCCP with our mitochondrial preparation from both brain and muscle with the condition suggested by the literature [17] (Fig. 1). To optimize such measurement, we decide to modulate the concentrations of FCCP in the measurements with the optimal amounts of samples we just identified (Figs. 3 and 5). 1. Measure the OCR with FCCP at 1, 2, 3, 4, 5, 6, 7, and 8 μM using 2 μg Brain samples (see Note 10). 2. Measure the OCR with FCCP at 1, 2, 3, 4, 5, 6, 7, and 8 μM using 1 μg Muscle samples (see Note 10).

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Fig. 2 Determination of optimal amounts of brain mitochondrial preparation in the measurement of basal OCR induced by ADP. The coupling experiment is performed with Succinate (10 mM) as the substrate in the presence of Rotenone (2 μM) and ADP (4 mM) as described in methods. (a) OCR measurements with 0.3–7 μg samples, and is further divided into three groups as the sample amounts with 0.3–2 μg samples, the slope is 82.28 (a). 2–5 μg samples, the slope is 138.3 (b). 5–7 μg samples, the slope is 37.84 (c)

Overall, we find tissue-specific responses to ADP and FCCP induction in respiration in our system, which is in line with our previous finding on the specific regulation of respiratory machinery in brain and muscle [16, 18]. Importantly, we identify the optimal conditions for such measurements which are not always in line with the recommendations from the previous studies.

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Fig. 3 Determination of optimal amounts of brain mitochondrial preparations in FCCP induced respiration. The FCCP induced OCR is measured after recording both basal and uncoupled respiration, with subsequent addition of FCCP as shown in Fig. 1. (a) OCR measurements with 0.3–7 μg samples, and is further divided into three groups as the sample amounts with 0.3–2 μg samples, the slope is 76.39 (a). 2–5 μg samples, the slope is 118.6 (b). 5–7 μg samples, the slope is 21.26 (c)

Acknowledgments This work is supported by grants from National Institute of Health (R01 GM109434 and GM130129), the Pathology Core in the San Antonio Nathan Shock Center (P30-AG013319), and LT and YB are also supported by William and Ella Owens Medical Research Foundation.

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Fig. 4 Determination of optimal amounts of brain mitochondrial preparation in the measurement of basal OCR induced by ADP. The coupling experiment is performed with Succinate (10 mM) as the substrate in the presence of Rotenone (2 μM) and ADP (4 mM) as described in methods. (a) OCR measurements with 0.2–7 μg samples, and is further divided into three groups as the sample amounts with 0.2–0.3 μg samples, the slope is 34.08 (a). 0.3–3 μg samples, the slope is 191.2 (b). 3–7 μg samples, the slope is 23.85 (c)

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Fig. 5 Determination of optimal amounts of muscle mitochondrial preparations in FCCP induced respiration. The FCCP induced OCR is measured after recording both basal and uncoupled respiration and treatment of FCCP as shown in Fig. 1. (a) OCR measurements with 0.2–7 μg samples, and is further divided into three groups as the sample amounts with 0.2–0.3 μg samples, the slope is 49.97 (a). 0.3–3 μg samples, the slope is 158.9 (b). 3–7 μg samples, the slope is 3.865 (c)

Fig. 6 Determination optimal FCCP concentration. Titration of FCCP (1–8 μM) with the optimal amounts of samples preparations. (a) 2 μg brain samples are used with 1, 2, 3, 4, 5, 6, 7, and 8 μM FCCP. (b) 1 μg Muscle samples are used with 1, 2, 3, 4, 5, 6, 7, and 8 μM FCCP

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References 1. Bhatti JS, Bhatti GK, Reddy PH (2017) Mitochondrial dysfunction and oxidative stress in metabolic disorders — a step towards mitochondria based therapeutic strategies. Biochim Biophys Acta Mol Basis Dis 1863(5): 1066–1077. https://doi.org/10.1016/j. bbadis.2016.11.010 2. Vartak R, Porras CA-M, Bai Y (2013) Respiratory supercomplexes: structure, function and assembly. Protein Cell 4(8):582–590. https:// doi.org/10.1007/s13238-013-3032-y 3. Ramsay RJC (2019) Electron carriers and energy conservation in mitochondrial respiration. 5:1–14 4. Porras CA, Bai Y (2015) Respiratory supercomplexes: plasticity and implications. Front Biosci (Landmark Ed) 20:621–634 5. Haas RH (2019) Mitochondrial dysfunction in aging and diseases of aging. Biology (Basel) 8 ( 2 ) : 4 8 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / biology8020048 6. Ghezzi D, Zeviani M (2018) Human diseases associated with defects in assembly of OXPHOS complexes. Essays Biochem 62(3): 2 7 1 – 2 8 6 . h t t p s : // d o i . o r g / 1 0 . 1 0 4 2 / EBC20170099 7. Sharma LK, Lu J, Bai Y (2009) Mitochondrial respiratory complex I: structure, function and implication in human diseases. Curr Med Chem 16(10):1266–1277. https://doi.org/ 10.2174/092986709787846578 8. Tegelberg S, Tomasˇic´ N, Kallij€arvi J, Purhonen J, Elme´r E, Lindberg E, Nord DG, Soller M, Lesko N, Wedell A, Bruhn H, Freyer C, Stranneheim H, Wibom R, Nennesmo I, Wredenberg A, Eklund EA, Fellman V (2017) Respiratory chain complex III deficiency due to mutated BCS1L: a novel phenotype with encephalomyopathy, partially phenocopied in a Bcs1l mutant mouse model. Orphanet J Rare Dis 12(1):73. https://doi. org/10.1186/s13023-017-0624-2 9. Grossman LI, Shoubridge EA (1996) Mitochondrial genetics and human disease. BioEssays 18(12):983–991. https://doi.org/10. 1002/bies.950181208 10. Wallace DC (2001) Mouse models for mitochondrial disease. Am J Med Genet 106(1): 71–93. https://doi.org/10.1002/ajmg.1393

11. Melov S, Coskun PE, Wallace DC (1999) Mouse models of mitochondrial disease, oxidative stress, and senescence. Mutat Res 434(3): 233–242. https://doi.org/10.1016/s09218777(99)00031-2 12. Bahr T, Welburn K, Donnelly J, Bai Y (2020) Emerging model systems and treatment approaches for Leber’s hereditary optic neuropathy: challenges and opportunities. Biochim Biophys Acta Mol Basis Dis 1866(6): 165743. https://doi.org/10.1016/j.bbadis. 2020.165743 13. Kruse SE, Watt WC, Marcinek DJ, Kapur RP, Schenkman KA, Palmiter RD (2008) Mice with mitochondrial complex I deficiency develop a fatal encephalomyopathy. Cell Metab 7(4):312–320. https://doi.org/10. 1016/j.cmet.2008.02.004 14. Hofhaus G, Shakeley RM, Attardi G (1996) Use of polarography to detect respiration defects in cell cultures. Methods Enzymol 264:476–483. https://doi.org/10.1016/ S0076-6879(96)64043-9 15. Bai Y, Attardi G (1998) The mtDNA-encoded ND6 subunit of mitochondrial NADH dehydrogenase is essential for the assembly of the membrane arm and the respiratory function of the enzyme. EMBO J 17(16):4848–4858. https://doi.org/10.1093/emboj/17.16. 4848 16. Li H, Kumar Sharma L, Li Y, Hu P, Idowu A, Liu D, Lu J, Bai Y (2013) Comparative bioenergetic study of neuronal and muscle mitochondria during aging. Free Radic Biol Med 63:30–40. https://doi.org/10.1016/j.fre eradbiomed.2013.04.030 17. Rogers GW, Brand MD, Petrosyan S, Ashok D, Elorza AA, Ferrick DA, Murphy AN (2011) High throughput microplate respiratory measurements using minimal quantities of isolated mitochondria. PLoS One 6(7):e21746. https://doi.org/10.1371/journal.pone. 0021746 18. Li Y, Li H-Z, Hu P, Deng J, Banoei MM, Sharma LK, Bai Y (2009) Generation and bioenergetic analysis of cybrids containing mitochondrial DNA from mouse skeletal muscle during aging. Nucleic Acids Res 38(6): 1913–1921. https://doi.org/10.1093/nar/ gkp1162

Chapter 9 Simultaneous Acquisition of Mitochondrial Calcium Retention Capacity and Swelling to Measure Permeability Transition Sensitivity Arielys M. Mendoza and Jason Karch Abstract The loss of mitochondrial cristae integrity and mitochondrial swelling are hallmarks of multiple forms of necrotic cell death. One of the most well-studied and relevant inducers of mitochondrial swelling is matrix calcium (Ca2+). Respiring mitochondria will intake available Ca2+ into their matrix until a threshold is reached which triggers the opening of the mitochondrial permeability transition pore (MPTP). Upon opening of the pore, mitochondrial membrane potential dissipates and the mitochondria begin to swell, rendering them dysfunctional. The total amount of Ca2+ taken up by a mitochondrion prior to the engagement of the MPTP is referred to as mitochondrial Ca2+ retention capacity (CRC). The CRC/swelling assay is a useful tool for observing the dose-dependent event of mitochondrial dysfunction in real-time. In this technique, isolated mitochondria are treated with specific boluses of Ca2+ until they reach CRC and undergo swelling. A fluorometer is utilized to detect an increase in transmitted light passing through the sample as the mitochondria lose cristae density, and simultaneously measures calcium uptake by way of a Ca2+-specific membrane impermeable fluorescent dye. Here we provide a detailed protocol describing the mitochondrial CRC/swelling assay and we discuss how varying amounts of mitochondria and Ca2+ added to the system affect the dose-dependency of the assay. We also report how to validate the assay by using MPTP and calcium uptake inhibitors and troubleshooting common mistakes that occur with this approach. Key words Mitochondria, Fluorometry, Calcium retention capacity, CRC, Calcium Green 5 N, Mitochondrial dysfunction, Cell death, Mitochondrial swelling, Mitochondrial permeability transition pore (MPTP)

1

Introduction The disruption of calcium homeostasis plays a significant role in numerous pathological events such as ischemic injuries and degenerative disorders [1]. During such states, cytosolic calcium (Ca2+) levels increase above physiological thresholds, which then influx into the matrix of mitochondria [2]. High concentration of mitochondrial matrix Ca2+ triggers the opening of the mitochondrial permeability transition pore (MPTP), a nonspecific pore that

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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results in the permeabilization of the inner mitochondrial membrane and allows for the free flow of solutes up to 1.5 kDa in size [3]. Sustained MPTP opening induces complete mitochondrial dysfunction, characterized by loss of membrane potential, subsequent inhibition of ATP production, and mitochondrial swelling [4, 5]. Mitochondrial dysfunction is a hallmark of MPTP-dependent necrosis, which is a critical regulator of ischemic injuries, degenerative disorders, aging, and many other necrotic death related diseases [6–9]. Mitochondrial function is often used as a marker of cellular viability. Inversely, mitochondrial dysfunction serves as a marker of cellular damage and can be measured in several ways; morphological changes by transmission electron microscopy (TEM), loss of membrane potential by membrane-potential-dependent fluorescent dyes, decreases in mitochondrial respiration or capacity by measuring oxygen consumption, or by analyzing the accumulations of mitochondrial DNA mutations [10–12]. Here we describe another powerful technique to assess the sensitivity of Ca2+-dependent mitochondrial dysfunction in real time, known as the calcium retention capacity (CRC) assay and the mitochondrial swelling assay. CRC assay is an ex vivo fluorometric method performed by kinetically acquiring changes in fluorescence when mitochondria are challenged with Ca2+ [13]. The mitochondrial swelling assay is an ex vivo light absorbance method performed by kinetically acquiring changes in transmitted light following Ca2+ stimulation. Fortunately, a fluorometer that is equipped with two detectors, one to acquire fluorescence and the other to acquire transmittance, can simultaneously obtain both parameters streamlining experimentation and increasing the efficiency per sample. The machine we use is the Photon Technology International (PTI) QuantaMaster 8000 Series from HORIBA Scientific where the detector acquiring fluorescence is positioned 90
from the sample and the light source, while the detector acquiring transmittance is positioned 180
from the sample and the light source. We utilize a cell-impermeable Ca2+ indicator Calcium Green™-5 N (CG5N) which becomes excited at a wavelength of 506 nm and emits fluorescence at 532 nm [14]. Upon transiently binding Ca2+ upon stimulation with Ca2+ the dye in the cuvette immediately fluoresces represented by an immediate peak [15]. Over time, mitochondria within the cuvette begin taking up the Ca2+, which is represented by a decrease in fluorescence (Fig. 1a, b). The mitochondria are continuously incubated with additional Ca2+ and the process is repeated until maximum Ca2+ capacity is achieved. At this point, the MPTP engages and the mitochondria swell and begin to extrude Ca2+ [16]. Simultaneously, the mitochondria begin to swell, which is exhibited by a continuous increase in transmitted light passing through the sample (Fig. 1b, c). In addition, CRC traces are quantified by taking

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Fig. 1 Schematic representation of mitochondrial CRC/swelling assay with representative traces. (a) Schematic representing the events within the cuvette during the mitochondrial CRC/swelling assay. Numbers coordinate to events displayed on the traces. (b) and (c) 2 mg of mitochondria continuously dosed with 20 μM Ca2+ (arrows) over specified increments of time until CRC and swelling events occur. Excitement of calcium green 5 N (CG5N) is shown by sharp increase in fluorescence at the time of calcium treatment (event 1). Mitochondrial uptake is represented by a subsequent decrease in fluorescence, demonstrating that Ca2+ becomes unbound from the dye and enters the mitochondrial matrix (event 2). CRC is achieved once there is a release of Ca2+ from the mitochondria demonstrated by the steady increase and eventual plateau in fluorescence (event 3). Mitochondrial swelling is indicated by the sharp decrease in absorbance, which represents the opening of the MPTP

into account the amount of Ca2+ injected into the cuvette versus the amount taken up by the mitochondria (Fig. 2). Notably, the quantitation of this assay relies heavily on the total number of mitochondria put into the cuvette as well as the concentration of each Ca2+ injection (Fig. 3). The combination of the swelling assay with the CRC assay not only doubles the amount of data acquired per analysis, but also increases the ability to determine the efficacy of pharmacological and/or genetic inhibitors and inducers of the MPTP. Compounds that are commonly used to validate the CRC and swelling assays include the cyclophilin D (CypD) inhibitor, cyclosporin A (CsA), and the substrate of the adenine nucleotide translocator (ANT), adenosine diphosphate (ADP), or the mitochondrial calcium uniporter (MCU) inhibitor, Ru360, which is an oxygen-bridged di-nuclear ruthenium amine complex. Both CRC and

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mitochondrial swelling rely on mitochondrial Ca2+ uptake through the MCU, therefore, the inhibition of the MCU by Ru360 prevents calcium uptake and consequentially, prevents mitochondrial swelling [17, 18]. Thus, Ru360 is used to validate the CRC assay’s ability to measure mitochondrial Ca2+ uptake, and when added to the sample, a stair-step pattern in fluorescence as well as no change in absorbance is observed following multiple additions of Ca2+ (Fig. 4). To increase mitochondrial CRC, CsA, and/or ADP can be added to the sample [19]. When individually added, CsA- or ADP-treated mitochondria can intake significantly more Ca2+ before the MPTP engages and swelling occurs (Fig. 5a, b). However, in combination, mitochondria become highly resistant to Ca2+-induced MPTP opening, which results in a further increase in CRC (Fig. 5). The origins of the mitochondrial swelling assay began with experiments in the mid-twentieth century, which were conducted using standard spectrophotometry [20–22]. Due to the advancement of florescent dyes and methods of detection, the classic mitochondrial swelling assay has evolved into the modern CRC/ swelling assay, enabling researchers to examine Ca2+ uptake into mitochondria prior to mitochondrial swelling and increasing the amount of data obtained. In addition to swelling, total Ca2+ capacity and the rate of Ca2+ uptake may now be obtained concurrently. The combination of these analyses into a single assay is an extremely powerful tool to study MPTP sensitivity and mitochondrial dysfunction.

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2

Materials All the following solutions and isolated biological samples should remain at 4
C or on ice, except for the assay buffer, which can remain at room temperature.

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1. Isolation buffer: 225 mM mannitol, 75 mM sucrose, 5 mM HEPES, 1 mM EGTA, pH 7.4. Add 20.5 g mannitol, 12.8 g sucrose, 0.60 g HEPES, and 1 mL of 0.5 M EGTA into a 1 L beaker. Add 400 mL water and stir until dissolved, cool overnight at 4
C. Next day, adjust pH with KOH and HCl and bring up the final volume to 500 mL. Store at 4
C. 2. Assay buffer: 125 mM KCl, 20 mM HEPES, 2 mM KH2PO4, 40 μM EGTA, pH to 7.2. Weigh 4.7 g KCl, 2.8 g HEPES, 0.14 KH2PO4 in a 500 mL beaker. Add 0.04 mL of 0.5 M EGTA and raise volume up to 400 mL with water while stirring. Adjust pH using KOH and HCL to 7.2 and raise the final volume to 500 mL with additional water. Store at room temperature. 3. 8 mL glass homogenizer, with polytetrafluoroethylene (PTFE) pestle. 4. Standard temperature controlled large centrifuge with a fixed rotor capable of housing 15 mL volume conical tubes.

2.2 CRC Assay Reagents and Equipment

1. Calcium Green™-5 N (CG5N): 50 μM stock concentration. Add 8.39 mL water to 500 μg, aliquot and store at 20
C within a desiccator. Keep from light. 2. Sodium pyruvate: 1 M stock concentration, pH 7.2. Add 3.3 g of pyruvate into 30 mL of assay buffer. Aliquot (1 mL) and store at 20
C. 3. DL-Malic acid: 1 M stock concentration, pH 7.2. Add 4 g of DL-Malic acid into 30 mL assay buffer. Aliquot (1 mL) and store at 20
C. 4. Calcium Chloride (CaCl2, Ca2+): 1 M stock concentration. Dissolve 3.3 g of CaCl2 into 30 mL deionized water (mild exothermic reaction). To make 10 mM CaCl2, add 120 μL of 1 M CaCl2 into 12 mL final volume of deionized water. Vortex well and store at room temperature. 5. Cyclosporin A (CsA): 5 mM stock concentration. Dissolve 10 mg of CsA in 1.6 mL 100% ethanol, aliquot and store at 20
C. 6. Adenine Diphosphate (ADP): 50 mM stock concentration. Dissolve 5 mg ADP in 585 mL of assay buffer, aliquot and store at 20
C. 7. Ru360: 5 mM stock concentration. Dissolve 500 μg Ru360 in 182.9 μl of deionized water. Aliquot and store protected from light at 20
C. 8. Fluorometer (Photon Technology International (PTI) QuantaMaster 8000 Series, Horiba Scientific). 9. Photon Technology International (PTI) FelixGX software. 10. 3500 μL macro fluorescent cuvette, glass quartz.

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11. Small cuvette stool square for elevating the 1 mL sample into the path of the laser. 12. 1.5 mm flea micro stir bar. 13. Laboratory grade swabs.

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Methods

3.1 Mitochondrial Isolation from Mouse Liver

Cool the centrifuge to 4
C prior to mitochondrial isolation. 1. Euthanize mouse and carefully access abdominal cavity. 2. Dissect largest lobe of liver from mice. In a weigh boat, wash with mitochondrial isolation buffer. 3. Transfer tissue to a small beaker (50 mL) containing 7 mL of ice cold isolation buffer. Keep on ice. 4. While on ice, finely mince the lobe with scissors (1–2 mm pieces) and transfer to 8 mL glass tissue homogenizer (see Note 1). 5. Dounce with a PTFE pestle until the mixture is homogenous and all pieces of tissue are no longer visible (7–10 plunges) (see Note 2) and transfer into a 15 mL conical. 6. Centrifuge sample at 800  g for 5 min at 4
C. 7. Remove sample from centrifuge. Transfer supernatant into new 15 mL conical. Discard pellet. 8. Centrifuge sample at 10,000  g for 10 min at 4
C. 9. Remove sample from centrifuge. Aspirate the supernatant, careful not to aspirate the pellet. 10. While on ice, reconstitute the pellet in 7 mL of chilled isolation buffer by gently pipetting up and down until the pellet is suspended. 11. Centrifuge for an additional 10 min at 10,000  g at 4
C. 12. Aspirate supernatant and completely suspend pellet in 1 mL of assay buffer. Transfer into 1.7 mL micro-centrifuge tube. Keep on ice. 13. Determine protein concentration of the sample. Record concentration in μg/μL and keep on ice (see Note 3).

3.2 CRC/ Swelling Assay

1. Prepare required equipment and reagents (see Notes 4 and 5). 2. Load 2 mg of mitochondria into the cuvette per assay. Calculate the volume per assay necessary based on determined sample concentration (see Note 6). 3. Calculate the volume of reagents and mitochondria necessary per run and add them into a clean quartz cuvette with the stir bar. Final concentrations of reagents in the cuvette containing

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1 mL final volume: 7 mM pyruvate, 1 mM malic acid, 0.5 μM CG5N, and lastly add the 2 mg of mitochondria (see Notes 7 and 8). 4. Once all components are added into the cuvette, place cuvette into the fluorometer device and incubate for 5 min with stirring with the lid closed. 5. Click “start” on the FelixGX™ program to obtain a baseline measurement. 6. When adding Ca2+ to the cuvette, press the “pause” button, open the lid of the fluorimeter and manually add Ca2+ (2 μL of 10 mM CaCl2 stock or other desired concentration) at a preferred time. Quickly close the lid and press “continue” to resume the assay (see Notes 9 and 10). 7. Once fluorescence has plateaued, add another dose of Ca2+ into the cuvette. 8. Repeat step 7 until mitochondrial swelling is observed, which is identified as a sharp and steady increase in transmittance (see Note 11). 9. Allow fluorescence and transmittance curve to plateau (time may vary dependent on the presence of inhibitors) and the press “stop” to end run. 10. Clean cuvette and stir bar thoroughly with swabs and deionized water. Prepare the desired following assay with additional inhibitors/alternate inducers of mitochondrial swelling (see Notes 12 and 13). 3.3 Data Analysis and Quantification

1. Save file on program and export all traces into a text (.TXT) file. The data can then be copied and pasted into another program such as Microsoft® Excel® for graphing and data analysis. 2. Convert transmittance values to absorbance using a baseline value and the
Beer–Lambert equa. tion T ¼ 2  log transmittance baseline 3. Mitochondrial swelling curves can be graphed as raw values. However, if there are large baseline differences between samples, graphing the traces relative to the baseline absorbance may be appropriate. 4. The fluorescent curves should be graphed using raw values since minor differences in baseline fluorescence is negligible to assay interpretation. 5. In order to quantify mitochondrial CRC, first determine the “total amount of Ca2+” added to the assay. This can be easily determined by counting the number of peaks in the graphs, total volume of Ca2+ added per peak, and knowing the CaCl2 stock used.

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6. Calculate the amount of “free Ca2+” (extra-mitochondrial Ca2+) at the point of MPTP opening (see Note 14). 7. Calculate CRC using the following equation: CRC ¼ ðtotal Ca2þfree Ca2þÞ (see Note 15). total mitochondria 8. Once sufficient experimental replicates are achieved to perform statistical analyses, the data can be represented qualitatively using a representative traces and quantitatively using the calculated CRC values (Fig. 2).

4

Notes 1. To help transfer all minced tissue at once efficiently, swirl beaker and quickly pour contents into homogenizer. Letting tissue settle at the bottom of the homogenizer, return some buffer into the beaker and repeat this method to transfer all remnants of tissue. 2. Gently homogenize tissue without forming bubbles and twist plunger during upward strokes to homogenize the tissue. 3. Invert micro-centrifuge tube containing sample to keep mitochondria from settling in bottom of the tube for a more accurate reading of protein concentration. Keep mitochondria on ice throughout the rest of the procedure. However, the mitochondria are also time-sensitive and will remain useful for the CRC/swelling assay for approximately 6 h. 4. Allow pyruvate, malic acid, and MPTP inhibitors such as cyclosporin A and ADP to defrost on ice until thawed. 5. Turn on the fluorometer for approximately 15 min prior to using to allow the light source to warm up completely. Connect to energy source, hit laser switch, press ignite button, and check for lamp light which should be green. Check to see if motor driver stirrer is on (should make a faint sound). 6. The CRC/mitochondrial swelling assay is very dosedependent, meaning the timing of achieving CRC and swelling is very dependent on the amount of mitochondria used, or the amount of calcium used per spike (Fig. 3). Keep this in mind to determine the optimal amount of either to use, if seeking to detect more sensitive analysis a greater amount of mitochondria and lower concentration of calcium is recommended. 7. The addition of 10 μM Ru360 into the cuvette prior to any additions of Ca2+ may be used to confirm if the assay is truly detecting mitochondrial Ca2+ uptake (Fig. 4). 8. The addition of 2 μM CsA and/or 300 μM ADP into to the cuvette prior to any additions of Ca2+ may be used as a positive control for MPTP desensitization (Fig. 5).

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9. Once the assay is resumed after adding Ca2+, the fluorescence curve should spike upwards immediately. First addition(s) to the cuvette will be buffered by the EGTA within the assay buffer (depending on the concentration of Ca2+ added), but subsequent additions will stimulate the response of the CG5N dye. 10. Do not open the lid of the fluorometer prior to pressing the “pause” button. Opening the lid will prematurely disrupt the assay and all recorded fluorescence and transmittance traces will drop to zero due to a shutter automatically blocking the light source when the lid is open. 11. Do not add additional calcium once CRC is reached and mitochondrial swelling begins (simultaneous event). This moment in the assay is evident by an increase followed by a plateau in fluorescent light and by a simultaneous increase in transmitted light. 12. The use of laboratory grade swabs to clean the cuvettes with water may help prevent residual inhibitors from affecting the following assays. 13. When performing multiple assays that will be qualitatively compared, use consistent time points when adding Ca2+ into the cuvette and allow all runs to transpire until MPTP opening occurs. 14. In order to calculate “free Ca2+,” the user must first calibrate the Calcium green 5 N dye on their fluorescent system. Once calibrated, the “free Ca2+” concentration at the point of MPTP opening can be determined by the fluorescent value at that time (Fig. 2). 15. Final CRC data mitochondria.”

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Acknowledgments This work was supported by the National Heart, Lung, and Blood Institute of the National Institutes of Health under award number R01HL150031 and the NIH training grant award number 5T32HL007676-29. References 1. Bernardi P, Lisa FD (2015) The mitochondrial permeability transition pore: molecular nature and role as a target in cardioprotection. J Mol Cell Cardiol 78:100–106 2. Gunter TE, Buntinas L, Sparagna G et al (2000) Mitochondrial calcium transport:

mechanisms and functions. Cell Calcium 28(5–6):285–296 3. Briston T, Roberts M, Lewis S et al (2017) Mitochondrial permeability transition pore: sensitivity to opening and mechanistic

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dependence on substrate availability. Sci Rep 7: 10492 4. Karch J, Kwong JQ, Burr AR et al (2013) Bax and Bak function as the outer membrane component of the mitochondrial transition pore. PNAS 111(29):10396–10397 5. Kroemer G, Reed J (2000) Mitochondrial control of cell death. Nat Med 6:513–519 6. Patel P, Karch J (2019) Regulation of cell death in the cardiovascular system. Int Rev Cell Mol Biol 353:153–209 7. Millay DP, Sargent MA, Osinska H et al (2008) Genetic and pharmacologic inhibition of mitochondrial-dependent necrosis attenuates muscular dystrophy. Nat Med 14(4):442–447 8. Panel M, Ghaleh B, Morin D (2018) Mitochondria and aging: a role for the mitochondrial transition pore? Aging Cell 17(4):e12793 9. Readnower RD, Hubbard WB, Kalimon OJ et al (2021) Genetic approach to elucidate the role of Cyclophilin D in traumatic brain injury pathology. Cell 10(2):199 10. Burattini S, Falcieri E (2013) Analysis of cell death by electron microscopy. Methods Mol Biol 1004:77–89 11. Crowley LC, Christensen ME, Waterhouse NJ (2016) Measuring mitochondrial transmembrane potential by TMRE staining. Cold Spring Harb Protoc 2016:pdb.prot087361 12. Martin DB, David GN (2011) Assessing mitochondrial dysfunction in cells. Biochem J 435(2):297–312 13. Harisseh R, Abrial M, Chiari P et al (2019) A modified calcium retention capacity assay clarifies the roles of extra-and intracellular calcium pool in mitochondrial permeability transition pore opening. J Biol Chem 294:15282–15292 14. Rajdev S, Reynolds IJ (1993) Calcium green5N, a novel fluorescent probe for monitoring

high intracellular free Ca2+ concentrations associated with glutamate excitotoxicity in cultured rat brain neurons. Neurosci Lett 162(1–2):149–152 15. Deak AT, Jean-Quartier C, Bondarenko AI et al (2015) Assessment of mitochondrial Ca2+ uptake. Methods Mol Biol 1264:421– 439 16. Javadov S, Chapa-Dubocq X, Makarov V (2018) Different approaches to modeling analysis of mitochondrial swelling. Mitochondrion 38:58–70 17. Marchi S, Pinton P (2014) The mitochondrial calcium uniporter complex: molecular components, structure and physiopathological implications. J Physiol 592(5):829–839 18. Pan X, Liu J, Nguyen T, Liu C et al (2013) The physiological role of mitochondrial calcium revealed by mice lacking the mitochondrial calcium uniporter. Nat Cell Biol 15:1464–1472 19. Karch J, Bround MJ, Khalil H et al (2019) Inhibition of mitochondrial permeability transition by deletion of the ANT family and CypD. Science. Advances 5:EAAW4597 20. Tedeschi H, Harris DL (1955) The osmotic behavior and permeability to non-electrolytes of mitochondria. Arch Biochem Biophys 58: 52–57 21. Beavis AD, Brannan RD, Garlid KD (1985) Swelling and contraction of the mitochondrial matrix. I. A structural interpretation of the relationship between light scattering and matrix volume. J Biol Chem 260:13424– 13433 22. Selwyn MJ (1986) Use of transmittance to record mitochondrial swelling. Biochem Soc Trans 14(6):1045–1046

Chapter 10 Measuring Mitochondrial Function: From Organelle to Organism Matthew T. Lewis, Yan Levitsky, Jason N. Bazil, and Robert W. Wiseman Abstract Mitochondrial energy production is crucial for normal daily activities and maintenance of life. Herein, the logic and execution of two main classes of measurements are outlined to delineate mitochondrial function: ATP production and oxygen consumption. Aerobic ATP production is quantified by phosphorus magnetic resonance spectroscopy (31PMRS) in vivo in both human subjects and animal models using the same protocols and maintaining the same primary assumptions. Mitochondrial oxygen consumption is quantified by oxygen polarography and applied in isolated mitochondria, cultured cells, and permeabilized fibers derived from human or animal tissue biopsies. Traditionally, mitochondrial functional measures focus on maximal oxidative capacity—a flux rate that is rarely, if ever, observed outside of experimental conditions. Perhaps more physiologically relevant, both measurement classes herein focus on one principal design paradigm; submaximal mitochondrial fluxes generated by graded levels of ADP to map the function for ADP sensitivity. We propose this function defines the bioenergetic role that mitochondria fill within the myoplasm to sense and match ATP demands. Any deficit in this vital role for ATP homeostasis leads to symptoms often seen in cardiovascular and cardiopulmonary diseases, diabetes, and metabolic syndrome. Key words Aerobic metabolism, ADP sensitivity, Free energy homeostasis, Oxygen consumption, Magnetic resonance, Bioenergetics

1

Introduction Activities of daily living involve dynamic changes in energy demands that must be matched by energy production in the form of adenosine triphosphate (ATP). Failure to meet these demands results in a loss of energy homeostasis in the afflicted tissues and clinically presents with a broad spectrum of pathologies. For example, the loss of energy homeostasis in skeletal muscle results in elevated plasma lactate and creatine kinase, loss in cell viability, and exercise intolerance. Skeletal muscle ATP content would last less than 2 s during intense workloads in the absence of

Matthew T. Lewis and Yan Levitsky contributed equally to the writing of this manuscript. Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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replenishment by energy metabolism [1, 2], principally produced by mitochondrial oxidative phosphorylation. Thus, mitochondrial dysregulation or dysfunction is a potential underlying cause in many diseases where impaired exercise capacity is a presenting symptom [3–7]. While mitochondria contribute to redox homeostasis [8, 9], anaplerosis [10], and a number of other important biochemical processes within the cell [11], the sensing and matching of ATP demands through sensitivity to cytosolic ADP is their primary function [1, 3, 12]. Because of the importance of aerobic metabolism to our health and understanding of so many diseases mitochondria have been studied for over 70 years. However, many studies reporting mitochondrial dysfunction have primarily focused on maximal respiratory rates quantified by measuring oxygen consumption by the isolated organelle in the presence of saturating ADP concentrations. This is a useful measurement but is neither physiologic nor does it fully define mitochondrial function. Herein we describe how to challenge mitochondria across a broad range of physiological stimuli (e.g., submaximal ADP levels) including the maximal respiration rate. A plot of this function (mitochondrial respiration versus [ADP]) demonstrates ADP sensitivity, a more physiological descriptor of mitochondrial function. In the sections that follow, quantification of mitochondrial function including ADP sensitivity is outlined using a broad spectrum of measurements from the tissue level in vivo to cultured cells, permeabilized fibers, and isolated mitochondria in vitro. Herein, the measurements of mitochondrial function are separated into ATP production or oxygen consumption to quantify mitochondrial output or input functions, respectively (Fig. 1). Phosphorus magnetic resonance spectroscopy (31PMRS) quantifies ATP production across a range of workloads imposed on the muscle in vivo. This generates a range of [ADP] and permits definition of ADP sensitivity while operating at intrinsic ionic strength, pH, and oxygen tension. However, stimulating mitochondrial respiration toward maximal respiration in vivo is limited by muscle fatigue and glycolytic activation above 60% VO2max, factors which modulate the adenylate pool and complicate interpretation [13–15]. To circumvent this problem, purified mitochondrial preparations are used to query function independent of cytosolic complexities and enable mitochondria to achieve higher respiration rates than possible in vivo, generating useful information about detailed mitochondrial function but under specialized circumstances [12]. This strategy has since been applied in cell culture and permeabilized fibers to quantify mitochondrial function from maximal respiration rates. However, kinetic studies focused on ADP sensitivity in these preparations are not reflective of mitochondrial function due to diffusion limitations inherent to the preparation. We emphasize the importance of this concept by demonstrating “high” K0.5

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Fig. 1 Mitochondrial ATP production is necessary for prolonged exercise or maintaining activities of daily living. At the onset of activity, mitochondria are stimulated primarily by ADP to produce ATP and match demand. The capacity or function for mitochondria to sense and match a given demand is routinely assessed by in vitro (oxygen consumption) or in vivo (31PMRS) techniques. Together these techniques can be applied to determine the benefits of interventions such as exercise training or deficits invoked by disease

values for ADP sensitivity measurements in permeabilized fibers are a consequence of intra-fiber ADP diffusion gradients. To do this, we developed a simplified and concise reaction–diffusion model and recreated the permeabilized ADP sensitivity experiments in silico. In the model, we used the K0.5 value determined from isolated mitochondrial studies and were able to obtain “high” K0.5 ADP sensitivity values observed in the permeabilized fiber studies when correctly accounting for ADP diffusion and physiological ATP consumption rates.

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Materials

2.1 Phosphorus Magnetic Resonance Spectroscopy

1. MRS systems for routine clinical studies or dedicated research scanners can be used. The higher the field strength available, the better the signal to noise possible and hence higher temporal resolution. For example, higher field strengths (measured in Telsa) are required when the signal is low (e.g., small muscle mass). However, as magnet power increases, bore size are smaller because of the difficulty in maintaining field

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homogeneity. Thus, more powerful magnets are usually only used on able to study smaller subjects (e.g., rats and mice) while clinical MRI systems are used for human subjects. 2. Dual tuned (1H-31P) surface coil designed specifically for tissue being investigated operating in transmit and receive modes are most widely used. Surface coils generate a spherical field symmetric around the coil and thus penetrate the sample as a function of the radius so a 1 cm coil can project 0.5 cm into the sample. See Gadian [16] for a more detailed approach to coil design. 3. Magnet compatible ergometer and strain gage to quantify muscle work (e.g., force transducer). This is necessary to ensure the workload is maintained throughout the exercise protocol and is consistent at all intensities. If performing involuntary work, as with mammals under anesthesia, a muscle stimulator is used to pace the nerve or muscle directly. 4. Data analysis software must be able to import free induction decays and filter as needed. After data is filtered, software for identifying and quantifying spectral peaks such as jMRUi [17] is necessary. More software is listed and discussed in the consensus 31PMRS work by Meyerspeer et al. [18]. 2.2 Isolated Mitochondria

1. Sharp scissors. 2. Glass cannula. 3. Forceps. 4. Protease. 5. Tissue homogenizer. 6. Cardioplegia Buffer. (a) 24 mM KCl. (b) 100 mM NaCl. (c) 10 mM dextrose. (d) 25 mM MOPS. (e) 1 mM EGTA. (f) Titrated to pH 7.15. 7. Isolation Buffer. (a) 200 mM mannitol. (b) 50 mM sucrose. (c) 5 mM K2HPO4. (d) 0.1% (w/v) bovine serum albumin. (e) Titrated to pH 7.15. 8. Respiration buffer.

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(a) 130 mM KCl. (b) 5 mM K2HPO4. (c) 20 mM MOPS. (d) 1 mM MgCl2. (e) 0.1% (w/v) bovine serum albumin. (f) Titrated to pH 7.1 at 37
C. 9. Refrigerated centrifuge. 10. Substrates. (a) Pyruvate, malate, glutamate, succinate, glucose, and lactate. 11. Purified creatine kinase, creatine, phosphocreatine, ATP. 2.3 Mitochondrial Fractional Volume

1. Spectrophotometer, 1 mL cuvettes.

2.3.1 Cytochrome c Oxidase

3. Potassium ferracyanide.

2. ddH2O. 4. Sodium dithionite. 5. Assay Buffer. (a) 100 mM KH2PO4, pH 7.0. 6. 1 mM cytochrome c in 10 mM KH2PO4. 7. Homogenization Buffer. (a) 100 mM Tris-HCl. (b) 0.1% Triton X-100.

2.3.2 Citrate Synthase

1. Spectrophotometer, 1 mL cuvettes. 2. 1 M Tris-HCl pH 8.1. 3. Homogenization buffer. (a) 0.1 M Tris-HCl. (b) 0.005 M DTT. (c) 0.1% Triton X-100. 4. 1 mM DTNB in 1 M Tris-HCl pH 8.1. 5. 10 mM acetyl-CoA in ddH2O. 6. 10 mM oxaloacetate.

3

Methods

3.1 Phosphorus Magnetic Resonance Spectroscopy (31PMRS)

31

PMRS quantitatively measures metabolite concentrations within tissues including inorganic phosphate (Pi), phosphocreatine (PCr), and ATP. Utilizing the creatine kinase (CK) equilibrium, changes in key phosphate-containing metabolites under different

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Fig. 2 ATP free energy cycling in skeletal muscle. ATP is consumed in skeletal muscle to provide energy required for actin-myosin, SERCA, and Na+/K+ ATPases generating ADP and Pi. ATP supply matches demands through mitochondrial oxidative phosphorylation, glycolysis, and PCr buffering of ADP. Each of these systems communicate with the adenylate pool which is linked to the creatine kinase reaction allowing PCr to act as a reporter for cytosolic ADP levels which can be quantified by 31PMRS

perturbations permit quantification of muscle energetics and mitochondrial oxidative ATP production and their dynamic changes (Fig. 2). The CK equilibrium couples ATP production to ATP demand: ATPase : ATP ! ADP þ Pi þ αHþ CK equilibrium : ADP þ PCr þ Hþ $ ATP þ Cr Net : PCR þ βHþ ! Cr þ Pi where α (0.6) and β (0.4) represent partial protons produced as a function of environmental variables including temperature, pH, ionic strength, and pKa differences of substrates and products [19]. Using the CK equilibrium, oxidative phosphorylation can be monitored through changes in metabolites (PCr, Pi) during the transition from rest to a graded exercise state, during steadystate exercise, and in the transition back to rest immediately following exercise. The kinetics of PCr changes within the tissue are an important indicator of ATP and ADP dynamics and very useful biochemically. Specifically, the rate of PCr breakdown is proportional to the cellular ATPase rate and the rate of PCr recovery is directly related to the volume fraction of mitochondria within the tissue and used as a measure for tissue oxidative capacity [15, 20– 23]. The response to an energetic demand by skeletal muscle tissue is best described by the free energy of ATP hydrolysis, or phosphate potential. Free energy (kJ/mol) is calculated according to classical thermodynamics as follows:
 ½ADP½Pi ΔG ATP ¼ ΔG oATP þ 2:58  ln ½ATP

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Similarly, free ADP is calculated by rearrangement of the CK equilibrium: free ADP ¼

½ATP½Cr ½PCrK eq ½Hþ 

Quantifying free ADP and ATP with calculation of free energy during submaximal steady-state exercise bouts allows one to generate the mitochondrial transfer function, first proposed by Chance and colleagues [22]. The mitochondrial transfer function describes the rate of respiration for a given free energy or [ADP] and is one of the best indicators of mitochondrial respiratory function. Plotting mitochondrial oxidative phosphorylation (MOP) rate against [ADP] through a range of steady-state workloads defines the apparent ADP sensitivity, which is directly influenced by the volume of mitochondria within the tissue [24]. This means that while performing at equivalent workloads and oxygen consumption rates, [ADP] is higher in muscles with lower muscle mitochondrial content. Conversely, when mitochondrial content is doubled, the [ADP] is halved, reflected as increased ADP sensitivity [24–26]. Under certain circumstances ADP sensitivity cannot be interpreted as a direct measure of mitochondrial function. For example, if oxygen supply approaches the point where oxygen will limit respiration, ADP sensitivity is influenced  by the reduced oxygen QO2 2 supply to oxygen demand matching QO VO2 [3, 27]. In brief, as VO2 approaches a critical point, the influence of oxygen on ADP  sensi2 tivity starts to dominate the ADP function. Values of QO VO2 which lie near the critical point imply oxygen diffusion into the muscle cell has decreased. Under these conditions, ATP demand is maintained through metabolic adaptation (e.g., elevated [ADP] and glycolytic ATP production) whereas apparent ADP sensitivity is reduced. This can be observed in 31PMRS by demonstrating increased ADP for a given workload and increased PCr hydrolysis. Such results are evident in studies altering oxygen delivery during contractions without changing work rate [28]. Therefore, the investigator must know how close to the critical point they are working to account for oxygen limitations when interpreting results aimed at resolving mitochondrial dysfunction/dysregulation. This is critically important when working to understand disease conditions when the limits to respiration may not be well known. Confirming the lack of an oxygen limitations can be as simple as providing inspired gasses at 100% oxygen and showing no increase in respiration rate [29, 30]. 3.1.1 Protocols

1. Positioning subject: Whether human or animal, position the subject on gantry bed and place the NMR surface coil over the region of interest. Surface coils project at about a radius in

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depth into the sample and coil placement should be consistent between subjects due to muscle heterogeneity [31]. 2. Restraints for subject: Motion artifacts in magnetic resonance degrade the signal to noise (SNR) so subjects must be restrained, and for muscle experiments, the leg rigidly fixed. For humans this means padding and Velcro strapping and for animals more invasive methods can be used. For example, fixing the knee with a tungsten pin to reduce motion has been used in all our rodent studies [32, 33]. 3. RF filtering: All instrumentation monitoring the subject such as stimulation, force transducer, temperature, and respiration leads that run to the sample from external AC powered devices must be grounded to the magnet to reduce ground loops and radiofrequency noise. Leads also need to have a low pass RC or LC filter for each lead to eliminate antenna effects that degrade the signal-to-noise ratio (SNR). 4. Positioning in Magnet: Subjects are positioned in the magnet with the coil and adjacent sampled region as close to the magnet isocenter as possible to take advantage of the inherent magnet homogeneity. Electronic adjustments to the homogeneity (shimming) are easiest at the isocenter. 5. Tuning Receiver: The coil should be tuned in place to the frequency for phosphorus at the field strength being used. Tuning to the frequency is most important and matching to 50 Ohm impedance will ensure no reflected signal but this is often not perfect. The choice is always tune to frequency over matching. 6. Field Homogeneity: Homogeneity is accomplished automatically in modern MRI systems usually with gradient shimming protocols. Adjusting the field to account for local distortions due to the addition of the sample increases signal to noise for all nuclei including phosphorus. However, shimming can also be accomplished by using the available proton signal from the sample. Phosphorus coils even with very high Quality factors (Q) can still receive proton signal from sample water where the [1H] is 110 molar. 7. Temporal resolution: Where high time resolution is needed, signal acquisition is optimized by having a repetition time usually 10–20 times the instrument background OCR. (a) A final concentration of 0.5–2 million cells/mL in the sealed chamber is sufficient in a standard 2 mL Oroboros oxygraph with glucose, lactate, pyruvate as substrates. 7. Titrate oligomycin in 0.5–1 μM steps starting at 0.5 μM until no more response to addition is noted. Acquire 3–5 min of leak OCR. 8. Titrate FCCP (or CCCP) in 0.5–1.0 μM steps starting at 0.5 μM until a plateau or decrease is noted. Acquire 3–5 min of uncoupled OCR. 9. Add 10 μM rotenone + 10 μM antimycin A or 5 mM potassium cyanide. Acquire 3–5 min of OCRROX. See Notes 12 and 13 for specific details and suggestions for these protocols. 3.5 Permeabilized Fibers

For muscle cells, selective plasma membrane permeabilization with detergents (saponin) or bacterial toxins (streptolysin O) is often used to gain access to the cytosolic milieu [77, 93]. It is essential to run preliminary analyses to determine the optimal concentration of these permeabilizing agents to minimize non-specific effects [94]. However, because of the nature of this preparation, some important biophysical limitations preclude their use relative to kinetically driven questions such as ADP sensitivity. Retention of structurally organized organelles, cytoskeletal elements, and membrane bound proteins introduces complexities in assay design and data interpretation. Reagents in the bulk medium of permeabilized fiber preparations must traverse the intracellular space to reach target sites producing unavoidable diffusion limitations. Mass transport limitations will thus contribute to the observed kinetics and skew data collected using this system. Importantly, such mass transport phenomena are unlikely to play a role in the intact cell as ATP is hydrolyzed intracellularly, at the contractile apparatus, eliminating the longer distances of mass transport required in permeabilized fibers. In this section, we will demonstrate such effects using physiologically based computer simulations. The full suite of equations that govern the cellular processes under investigation are numerous and complex. That said, we can describe the system in simpler terms focusing on ADP diffusion and consumption by permeabilized fibers. Permeabilized fibers are an

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approximately 3000–5000 μm  70 μm cylindrical cluster of several myocytes [77, 95]. We will assume that permeabilized fibers are symmetrical along the radial axis and ignore diffusion along the longitudinal axis [96]. With these assumptions, the partial differential equation that describes ADP phosphorylation in the domain can be represented in cylindrical coordinates as follows: ! 2 ∂½ADP ∂ ½ADP 1 ∂½ADP ¼D  Rðr, ½ADPÞ: þ r ∂t ∂r 2 ∂r  Here, ∂½ADP is the time derivative of ADP along the domain ∂t from 0 to L where L is the radial distance (35 μm) from the surface of a permeabilized fiber to the center.2 D is the diffusion coefficient ½ADP for ADP equal to 150 μm2/s [47], ∂ ∂r is the local curvature of 2  the ADP concentration profile along the domain, and ∂½ADP is the ∂r local ADP concentration gradient along the domain. The term, R(r, [ADP]) represents an oxidative phosphorylation sink for ADP that may or may not depend on position along the domain. The partial differential equation above can be recast into an ordinary differential equation by solving it in the steady state as the following: 2

∂ ½ADP 1 ∂½ADP Rðr, ½ADPÞ þ : ¼ 2 r D ∂r ∂r If R is linear in [ADP], this equation has an analytical solution; however, when R is non-linear in [ADP], we must resort to numerical solvers. In this example, R is dependent on [ADP] in a non-linear, Michaelis–Menten like manner so that Rðr, ½ADPÞ ¼ ½ADP V max ½ADP þK 0:5 for all values of r between 0 and L. Here, Vmax is the maximum turnover of oxidative phosphorylation and K0.5 is the half-maximal saturation constant. In the below simulations, Vmax was set to 50 mM/min [95] and K0.5 was set to 50 μM [97]. With R defined this way, the above ordinary differential equation is written as 2

∂ ½ADP ½ADP 1 ∂½ADP ¼ : þχ r ½ADP þ K 0:5 ∂r 2 ∂r In this equation, χ equals Vmax/D so that when Vmax goes up or D goes down, [ADP] diffusion gradients along the domain are increased [98]. In contrast, when χ decreases, [ADP] diffusion gradients are flattened along the domain. Thus, the value of χ is a primary determinant of the [ADP] concentration profile along the domain and reflects changes to oxidative phosphorylation turnover capacity (e.g., mitochondrial density or metabolic state) as well as myocyte structure (e.g., ADP diffusion barriers). Before solving this equation, the boundary conditions must first be defined. For  the left boundary condition, we set the constraint as ∂½ADP  ∂r

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Fig. 7 (a) Simulated respiratory profiles as a function of buffer [ADP]o with calculated K0.5 values indicated for each χ. Respiration results were obtained by averaging JO2 values at each point along the domain r from 0 to L. (b) ADP concentration profiles along the domain at [ADP]o ¼ 1 mM. Colors represent different values of the χ, reflecting differences in metabolic activity of the fibers and degree of diffusional barriers present in the cytoplasm. Larger χ reflect higher metabolic activity of fibers and/or significant diffusional barriers. Small χ represent the inverse with Medium χ lying in the middle. JO2 was calculated from the rate of ADP phosphorylation assuming a P/O ratio of 2.7 [3]

ðr ¼ 0Þ ¼ 0, which is a no-flux boundary condition, based on the symmetry of the problem. For the right boundary condition, we enforce the constraint as [ADP](r ¼ L) ¼ [ADP]o where [ADP]o is the bulk medium concentration of ADP added to the permeabilized fiber suspension. With these two extra pieces of information, we solve the second order ordinary differential equation that governs the ADP concentration profile along the domain as a function of different values of χ that represent various conditions described above. Figure 7 shows the model simulation results after the above ordinary differential equation was solved in Matlab® using the bvp4c function. This function solves boundary value problems using the fourth-order method. Relative and absolute error tolerances were set to 105 and 108, respectively. These simulations reveal that significant ADP gradients form in myofibers which effectively pull the ADP sensitivity measure (K0.5) to the right. As shown in Fig. 7a, in the absence of diffusion barriers or low metabolic activity, ADP sensitivity approaches the value obtained from isolated mitochondria and 31PMRS in vivo studies [99]. As shown in Fig. 7b, increases in metabolic activity and/or diffusion restrictions produce heterogenous [ADP] profiles along r. At high χ, internal mitochondria are exposed to 5–10 lower steady state [ADP] compared to external mitochondria. Thus, the higher [ADP]o necessary to achieve maximum JO2 in permeabilized fibers reflects variable recruitment of internal mitochondria as opposed to saturation of binding sites, as assumed in more homogenous experimental models such as isolated mitochondria. While these simulations were generated using an idealized scenario, they demonstrate how permeabilized fiber studies can yield misleading ADP sensitivities lying in the 100 s of μM range.

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Such differences are observed in experimental models where the apparent K0.5 for ADP is tenfold greater in permeabilized muscle fibers compared to isolated mitochondria [99, 100]. Half-maximal respiration requires approximately twofold higher inorganic phosphate in permeabilized rat extensor digitorum longus (type-2) and soleus (type-1) compared to mitochondria isolated from the same tissue [101]. Likewise, oxygen affinities show similar patterns suggesting apparent K0.5 for oxygen of 50 μM in permeabilized fibers, significantly higher than the 0.1 μM estimate in isolated mitochondria [83, 102, 103]. While this complicates K0.5 interpretation relative to in vivo, changes in K0.5 between conditions as observed in diabetes [104, 105] or following bed rest [106] cannot be ignored. 3.6 Mitochondrial Fractional Volume

Irrespective of whether the preparations are in vivo muscles, in vitro isolated mitochondria, cells in culture, or permeabilized fibers all results require some aspect of normalization to the volume fraction of mitochondria present. In our hands there are several techniques for quantifying mitochondrial fractional volume. Mitochondrial enzyme activities have been correlated with content within tissue as quantified by electron microscopy to justify this approach [90]. For brevity, two of the most commonly used indicators of mitochondrial volume, cytochrome c oxidase, and citrate synthase activity, will be presented herein as abbreviated methods that are described in detail by Kasper [107]. These can be performed from any sample (e.g., cells, tissues, or mitochondria) that have been homogenized as described.

3.6.1 Cytochrome c Oxidase

1. Under fume hood, add a few grains of potassium ferricyanide to 10 mL ddH2O until dark yellow in color.

Step 1: Determine the Degree and Stability of Cytochrome c Reduction (% Reduction)

2. Produce a 50 μM cytochrome c solution in 10 mM KH2PO4 (pH 7.0 at 4
C), keep on ice. 3. Pipette 1 mL 50 μM cytochrome c solution into each of three cuvettes and record absorbance continuously at 550 nm at room temperature until a steady-state absorbance is recorded (~3 min). 4. Pipette 5 μL ferricyanide solution into each cuvette, mix, and continue recording. Ferricyanide will oxidize the cytochrome c causing a drop in absorbance. 5. Repeat to ensure cytochrome c is fully oxidized. 6. Record rate of absorbance change after complete oxidation reaches a steady state for ~3 min. If rate of reduction (increased absorbance) is significant, cytochrome c must be purified to avoid underestimation of activity quantified from muscle samples.

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7. Add a few grains of sodium dithionite to fully reduce the cytochrome c in solution. (a) Again, repeat this step to ensure complete reduction for accurate characterization of cytochrome c solution. 8. Calculate percent reduction of solution by comparing initial absorbance (step 3) to fully reduced (step 7). These should be approximately 90% reduced or greater. 9. Based on purity of the solution, re-calculate specific molecular weight for the concentration in solution. (a) This value will be used to determine quantity of solution for use in cytochrome c oxidase activity assays. Step 2: Quantify Cytochrome c Oxidase Activity Within Samples

1. Set spectrophotometer to 550 nm at 37
C. (a) Produce a 1 mM cytochrome c solution based on (step 1) in 10 mM KH2PO4. The final assay mixture will contain 60 μL of stock cytochrome c in a 1 mL reaction volume. 2. Using homogenization buffer, homogenize muscle samples at a 1:100 ratio to be assayed. 3. Load 900 μL assay buffer and 60 μL 1 mM cytochrome c solution per cuvette, mix by inversion. 4. Place cuvettes into spectrophotometer, allow to warm to 37
C for at (~5 min) with 1 mL of assay buffer as blank reference. 5. Record baseline absorbance for ~3 min. 6. Pipette 40 μL of 1:100 muscle homogenate sample per cuvette, mix, and continue recording. 7. Record reaction rate continuously for ~5 min. 8. Determine reaction rates (ABS/min) from the initial linear portion of reaction. Convert ABS/min to μmol cytochrome c oxidized per min using extinction coefficient of reduced cytochrome c (29,500 M1 cm1). 9. Normalize to gram muscle weight from 40 μL homogenate dilution.

3.6.2 Citrate Synthase

1. Set spectrophotometer to 412 nm and 37
C and keep all solutions on ice. 2. Pipette 900 μL assay reaction mixture (0.1 mM DTNB, 0.3 mM Acetyl-CoA, 0.1 M Tris-HCl pH 8.1) into cuvettes, place in spectrophotometer and allow to come up to temperature (~5 min). 3. Using homogenization buffer, homogenize muscle samples at a 1:100 ratio to be assayed.

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4. Add 50 μL 1:100 homogenate each to 3 cuvettes containing 900 μL assay reaction mixture. 5. Record background reaction rate for ~3 min. 6. Add 50 μL 10 mM Oxaloacetate to each cuvette to start the reaction, mix, and continue recording. 7. Record reaction rate continuously for ~5 min. 8. Determine reaction rates (ABS/min) from initial linear portion of reaction (+oxaloacetate) and subtract background reaction rate. 9. Normalize to gram muscle weight from 50 μL homogenate dilution.

4

Notes 1. Muscle contractions for many magnetic resonance experiments can be either isometric or isotonic and each have their own advantages/disadvantages in the restricted space of an MRI bore. Both generate a signal to stimulate oxidative metabolism but there are some limitations. Isotonic contractions require a range of motion and a resistive load which can be solved [108– 110] and quantified [111] but motion artifacts are a problem without gating contractions to data acquisition. For this reason isometric contractions are routinely used, especially in metabolic studies where metabolic efficiency is greater for this type of contraction [112]. 2. Ischemia has been used to measure basal ATPase rate or MOP from subsequent reperfusion and PCr recovery [39, 113, 114]. However, hypoxia complicates quantitation from non-oxidative contributions and potential activation of membrane ion pumps. 3. Isolation and respiration buffers. Isolation buffer: 200 mM mannitol, 50 mM sucrose, 5 mM K2HPO4, 5 mM MOPS, 1 mM EGTA at pH 7.15. Respiration buffer: 130 mM KCl, 5 mM K2HPO4, 20 mM MOPS, 1 mM MgCl2. Note, precise buffer composition depends on experimental aims or design. Chloride is known to lower ANT activity. As such, KCl can be replaced with equimolar K-lactobionate [72]. 4. An alternative to decapitation is a thoracotomy. When performing a thoracotomy to extract the heart, be sure the animal is heavily sedated and sprawled open by securing limbs with tape or some other means. Cut into the thorax at the sternum and cut up along the rib cage on both sides. After reaching the back sides, cut the ribs up toward the head and flip open the chest cavity. The heart is now exposed and ready for extraction.

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5. If mitochondrial purity is a primary concern (e.g., often the case with brain tissue), density gradient separation can be done in tandem with differential centrifugation [115–117]. 6. It is important to optimized substrate concentrations in respirometry assays [72]. While mitochondria from different tissue sources are more similar than they are different, there are key distinctions that require proper concentrations of substrate/ inhibitor combinations to ensure an unambiguous interpretation of the data. 7. The creatine kinase energetic clamp, as applied to respirometry, varies the of ATP hydrolysis, ΔG ATP ¼ ΔG oATP þ  free energy  ½ADP½Pi  , by modulating ½ATP ½ATP½Cr K eq ¼ ½PCr½ADP . Assuming

2:58 ln

the creatine kinase equilib-

rium, total phosphate changes as negligible, and a large and constant total creatine pool, the ½Cr free ADP concentration is recovered as: ½ADP ¼ ½KATP with eq ½PCr Keq ¼ 177 at 38
C, pH 7.0 and 1 mM Mg2+ [118].

8. Here we outlined a strategy to vary free [ADP], however, the SUIT titration is a popular method which yields the substratespecific leak state, ADP-stimulated (state 3), state 4o, uncoupled state (state 4u), and complex-IV specific turnover [83]. Residual OCR, after rotenone/antimycin A addition, corrects all states except complex-IV turnover which requires correction for TMPD auto-oxidation contained in the OCR after KCN titration. The apparent OCR for these states, barring complex IV turnover, is highly dependent on substrate mixtures provided for respiration. 9. SUIT protocols allow calculation of the respiratory control ratio (state 3:leak state) reflecting the relative intactness of mitochondrial preparations. 10. In the SUIT protocol maximal oxidative phosphorylation is attributed to state 3 OCR whereas maximal ETC turnover (state 3Eu) is typically attained after uncoupler titration and is almost always higher than state 3 [12]. The leak state is represented after substrate addition or after oligomycin addition to phosphorylating mitochondria. Presence of ATPases in the mitochondrial preparation may cause artifactual increases in apparent leak state without oligomycin if sufficient endogenous adenylates drive oxidative phosphorylation turnover. 11. As a measure of mitochondrial membrane integrity, the titration protocol can be modified by adding 10–20 μM of exogenous cytochrome c after ADP titration. Significant increases in OCR are diagnostic of outer mitochondrial membrane permeabilization and cytochrome c limited turnover, compromising interpretation of respiration relative to intact mitochondria.

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12. The whole cell respiratory control ratio (RCR) represents the attainable dynamic range of whole cell OCR. This measure is sensitive to inherent mitochondrial properties and extramitochondrial processes such as substrate uptake, metabolism, and transport. 13. Phosphorylation-linked OCR represents the portion of basal OCR dedicated to ATP generation for maintenance of cytosolic ΔGATP. This relation implies that increased inner membrane proton leak (OCRleak) results in decreased ATP generation through oxidative phosphorylation without concomitant increases in OCRbasal.

Acknowledgments This work was supported by financial support from the National Institutes of Health grants R00 HL121160, R01HL137694, R01 AG060731 and R01-DK095210 and fellowship F30EY030029. References 1. Meyer RA, Wiseman RW (2011) The metabolic systems: control of ATP synthesis in skeletal muscle. In: ACSM’s advanced exercise physiology, 2nd edn. Lippincott Williams & Wilkins, Philadelphia, pp 363–378 2. Hargreaves M, Spriet LL (2020) Skeletal muscle energy metabolism during exercise. Nat Metab 2:817–828 3. Lewis MT, Kasper JD, Bazil JN et al (2019) Quantification of mitochondrial oxidative phosphorylation in metabolic disease: application to type 2 diabetes. Int J Mol Sci 20(21): 5271 4. Weiss K, Sch€ar M, Panjrath GS et al (2017) Fatigability, exercise intolerance, and abnormal skeletal muscle energetics in heart failure. Circ Hear Fail 10(7):e004129 5. Hart CR, Layec G, Trinity JD et al (2018) Oxygen availability and skeletal muscle oxidative capacity in patients with peripheral artery disease: Implications from in vivo and in vitro assessments. Am J Physiol Heart Circ Physiol 315:H897–H909 6. Adami A, Cao R, Porszasz J et al (2017) Reproducibility of NIRS assessment of muscle oxidative capacity in smokers with and without COPD. Respir Physiol Neurobiol 235: 18–26 7. Genders AJ, Holloway GP, Bishop DJ (2020) Are alterations in skeletal muscle mitochondria a cause or consequence of insulin resistance? Int J Mol Sci 21:6948

8. Henriquez-Olguin C, Meneses-Valdes R, Jensen TE (2020) Compartmentalized muscle redox signals controlling exercise metabolism – current state, future challenges. 35: 101473 9. Murphy MP (2009) How mitochondria produce reactive oxygen species. Biochem J 417: 1–13 10. Balaban RS (2009) The role of Ca2+ signaling in the coordination of mitochondrial ATP production with cardiac work. Biochim Biophys Acta Bioenerg 1787(11):1334–1341 11. Brand MD (2014) The role of mitochondria in longevity and healthspan. Longev Heal 3:7 12. Brand MD, Nicholls DG (2011) Assessing mitochondrial dysfunction in cells. Biochem J 435:297–312 13. Brooks GA (1997) Importance of the “crossover” concept in exercise metabolism. Clin Exp Pharmacol Physiol 24:889–895 14. Forbes SC, Paganini AT, Slade JM et al (2009) Phosphocreatine recovery kinetics following low- and high-intensity exercise in human triceps surae and rat posterior hindlimb muscles. Am J Physiol Integr Comp Physiol 296:R161–R170 15. Paganini AT, Foley JM, Meyer RA (1997) Linear dependence of muscle phosphocreatine kinetics on oxidative capacity. Am J Physiol Cell Physiol 272(2 Pt 1):C501–C510

Mitochondria from In Vitro to In Vivo 16. Gadian DG (1982) Nuclear magnetic resonance and its applications to living systems. Oxford University Press, New York 17. Naressi A, Couturier C, Castang I et al (2001) Java-based graphical user interface for MRUI, a software package for quantitation of in vivo/ medical magnetic resonance spectroscopy signals. Comput Biol Med 31:269–286 18. Meyerspeer M, Boesch C, Cameron D et al (2020) P magnetic resonance spectroscopy in skeletal muscle: experts’ consensus recommendations. NMR Biomed 34(5):e4246 19. Meyer RA, Foley JM (1996) Cellular processes integrating the metabolic response to exercise. In: Comprehensive physiology. American Cancer Society, New York, pp 841–869 20. McCully KK, Fielding RA, Evans WJ et al (1993) Relationships between in vivo and in vitro measurements of metabolism in young and old human calf muscles. J Appl Physiol (1985) 75(2):813–819 21. Thompson CH, Kemp GJ, Sanderson AL et al (1995) Skeletal muscle mitochondrial function studied by kinetic analysis of postexercise phosphocreatine resynthesis. J Appl Physiol 78:2131–2139 22. Chance B, Leigh JS, Clark BJ et al (1985) Control of oxidative metabolism and oxygen delivery in human skeletal muscle: a steadystate analysis of the work/energy cost transfer function. Proc Natl Acad Sci 82:8384–8388 23. Kushmerick MJ, Meyer RA, Brown TR (1992) Regulation of oxygen consumption in fast- and slow-twitch muscle. Am J Physiol 263(3 Pt 1):C598–C606 24. Dudley GA, Tullson PC, Terjung RL (1987) Influence of mitochondrial content on the sensitivity of respiratory control. J Biol Chem 262:9109–9114 25. Holloszy JO (1967) Biochemical adaptations in muscle. Effects of exercise on mitochondrial oxygen uptake and respiratory enzyme activity in skeletal muscle. J Biol Chem 242: 2278–2282 26. Gollnick PD, Saltin B (1982) Significance of skeletal muscle oxidative enzyme enhancement with endurance training. Clin Physiol 2:1–12 27. Connett RJ, Honig CR, Gayeski TEJ et al (1990) Defining hypoxia: a systems view of V˙O2, glycolysis, energetics, and intracellular PO2. J Appl Physiol (1985) 68(3):833–842 28. Haseler LJ, Richardson RS, Videen JS et al (1998) Phosphocreatine hydrolysis during submaximal exercise: the effect of FI(O2). J Appl Physiol 85:1457–1463

169

29. Haseler LJ, Lin AP, Richardson RS (2004) Skeletal muscle oxidative metabolism in sedentary humans: 31P-MRS assessment of O2 supply and demand limitations. J Appl Physiol 97:1077–1081 30. Cree-Green M, Scalzo RL, Harrall K et al (2018) Supplemental oxygen improves in vivo mitochondrial oxidative phosphorylation flux in sedentary obese adults with type 2 diabetes. Diabetes 67:1369–1379 31. Boss A, Heskamp L, Breukels V et al (2018) Oxidative capacity varies along the length of healthy human tibialis anterior. J Physiol 596: 1467–1483 32. Kasper JD, Meyer RA, Beard DA et al (2019) Effects of altered pyruvate dehydrogenase activity on contracting skeletal muscle bioenergetics. Am J Physiol Integr Comp Physiol 316:R76–R86 33. Lewis MT, Kasper JD, Bazil JN et al (2019) Skeletal muscle energetics are compromised only during high-intensity contractions in the goto-kakizaki rat model of type 2 diabetes. Am J Physiol Regul Integr Comp Physiol 317: R356–R368 34. Harris RC, Hultman E, Nordesjo¨ LO (1974) Glycogen, glycolytic intermediates and highenergy phosphates determined in biopsy samples of musculus quadriceps femoris of man at rest. Methods and variance of values. Scand J Clin Lab Invest 33:109–120 35. Taylor DJ, Bore PJ, Styles P et al (1983) Bioenergetics of intact human muscle. A 31P nuclear magnetic resonance study. Mol Biol Med 1:77–94 36. Kemp GJ, Meyerspeer M, Moser E (2007) Absolute quantification of phosphorus metabolite concentrations in human musclein vivo by31P MRS: a quantitative review. NMR Biomed 20:555–565 37. Kushmerick MJ, Moerland TS, Wiseman RW (1992) Mammalian skeletal muscle fibers distinguished by contents of phosphocreatine, ATP, and Pi. Proc Natl Acad Sci U S A 89: 7521–7525 38. Adams GR, Foley JM, Meyer RA (1990) Muscle buffer capacity estimated from pH changes during rest-to-work transitions. J Appl Physiol 69:968–972 39. Blei ML, Conley KE, Kushmerick MJ (1993) Separate measures of ATP utilization and recovery in human skeletal muscle. J Physiol 465:203–222 40. Wiseman RW, Ellington WR, Rosanske RC (1989) Effects of extracellular pH and D-lactate efflux on regulation of intracellular pH during isotonic contractions in a

170

Matthew T. Lewis et al.

molluscan muscle: a 31p-nuclear magnetic resonance study. J Exp Zool 252:228–236 41. Blei ML, Conley KE, Odderson IR et al (1993) Individual variation in contractile cost and recovery in a human skeletal muscle. Proc Natl Acad Sci U S A 90:7396–7400 42. Foley JM, Meyer RA (1993) Energy cost of twitch and tetanic contractions of rat muscle estimated in situ by gated 31P NMR. NMR Biomed 6:32–38 43. Connett RJ, Gayeski TEJ, Honig CR (1985) Energy sources in fully aerobic rest-work transitions: a new role for glycolysis. Am J Physiol 248(6 Pt 2):H922–H929 44. Roman BB, Meyer RA, Wiseman RW (2002) Phosphocreatine kinetics at the onset of contractions in skeletal muscle of MM creatine kinase knockout mice. Am J Physiol Cell Physiol 283:C1776–C1783 45. Richardson RS, Wary C, Wray DW et al (2015) MRS evidence of adequate O2 supply in human skeletal muscle at the onset of exercise. Med Sci Sports Exerc 47:2299–2307 46. Meyer RA (1989) Linear dependence of muscle phosphocreatine kinetics on total creatine content. Am J Physiol Cell Physiol 257(6 Pt 1):C1149–C1157 47. Meyer RA, Sweeney HL, Kushmerick MJ (1984) A simple analysis of the “phosphocreatine shuttle”. Am J Physiol Cell Physiol 246(5 Pt 1):C365–C377 48. Gadian DG, Radda GK, Chance EM et al (1981) The activity of creatine kinase in frog skeletal muscle studied by saturation-transfer nuclear magnetic resonance. Biochem J 194(1):215–228 49. McGilvery RW, Murray TW (1974) Calculated equilibria of phosphocreatine and adenosine phosphates during utilization of high energy phosphate by muscle. J Biol Chem 249:5845–5850 50. Veech RL, Lawson JWR, Cornell NW et al (1979) Cytosolic phosphorylation potential. J Biol Chem 254(14):6538–6547 51. Gifford JR, Garten RS, Nelson AD et al (2016) Symmorphosis and skeletal muscle V˙O2 max: in vivo and in vitro measures reveal differing constraints in the exercise-trained and untrained human. J Physiol 594: 1741–1751 52. Andersen P, Saltin B (1985) Maximal perfusion of skeletal muscle in man. J Physiol 366: 233–249 53. Layec G, Malucelli E, Le FY et al (2013) Effects of exercise-induced intracellular acidosis on the phosphocreatine recovery kinetics: a 31P MRS study in three muscle groups in humans. NMR Biomed 26:1403–1411

54. Van Den BNMA, De FHMML, De GL et al (2007) Intersubject differences in the effect of acidosis on phosphocreatine recovery kinetics in muscle after exercise are due to differences in proton efflux rates. Am J Physiol Cell Physiol 293(1):C228–C237 55. Harkema SJ, Meyer RA (1997) Effect of acidosis on control of respiration in skeletal muscle. Am J Physiol Cell Physiol 272(2 Pt 1): C491–C500 56. Jubrias SA, Crowther GJ, Shankland EG et al (2003) Acidosis inhibits oxidative phosphorylation in contracting human skeletal muscle in vivo. J Physiol 553:589–599 57. Walter G, Vandenborne K, McCully KK et al (1997) Noninvasive measurement of phosphocreatine recovery kinetics in single human muscles. Am J Physiol Cell Physiol 272(2 Pt 1):C525–C534 58. Clark LC, Wolf R, Granger D et al (1953) Continuous recording of blood oxygen tensions by polarography. J Appl Physiol 6: 189–193 59. Levitsky Y, Pegouske DJ, Hammer SS et al (2019) Micro-respirometry of whole cells and isolated mitochondria. RSC Adv 9: 33257–33267 60. Gerencser AA, Neilson A, Choi SW et al (2009) Quantitative microplate-based respirometry with correction for oxygen diffusion. Anal Chem 81:6868–6878 61. Rogers GW, Brand MD, Petrosyan S et al (2011) High throughput microplate respiratory measurements using minimal quantities of isolated mitochondria. PLoS One 6: e21746 62. Divakaruni AS, Paradyse A, Ferrick DA et al (2014) Analysis and interpretation of microplate-based oxygen consumption and pH data. Elsevier, Amsterdam 63. Papkovsky DB, Dmitriev RI (2013) Biological detection by optical oxygen sensing. Chem Soc Rev. 42:8700–8732 64. Oomen PE, Skolimowski M, Verpoorte S (2016) Implementing oxygen control in chip-based cell and tissue culture systems. Lab Chip 16:3394–3414 65. Divakaruni AS, Wiley SE, Rogers GW et al (2013) Thiazolidinediones are acute, specific inhibitors of the mitochondrial pyruvate carrier. Proc Natl Acad Sci 110:5422–5427 66. Haller T, Ortner M, Gnaiger E (1994) A respirometer for investigating oxidative cell metabolism: toward optimization of respiratory studies. Anal Biochem 218:338–342 67. Duong QV, Hoffman A, Zhong K et al (2020) Calcium overload decreases net free

Mitochondria from In Vitro to In Vivo radical emission in cardiac mitochondria. Mitochondrion 51:126–139 68. Chance B, Williams GR (1955) Respiratory enzymes in oxidative phosphorylation. III. The steady state. J Biol Chem 217:409–427 69. Gnaiger E (2014) Mitochondrial pathways and respiratory control an introduction to OXPHOS analysis. Bioenerg Commun 2:122 70. Picard M, Ritchie D, Wright KJ et al (2010) Mitochondrial functional impairment with aging is exaggerated in isolated mitochondria compared to permeabilized myofibers. Aging Cell 9:1032–1046 71. Steinlechner-Maran R, Eberl T, Kunc M et al (1996) Oxygen dependence of respiration in coupled and uncoupled endothelial cells. Am J Physiol Cell Physiol 271(6 Pt 1): C2053–C2061 72. Wollenman LC, Vander PMR, Miller ML et al (2017) The effect of respiration buffer composition on mitochondrial metabolism and function. PLoS One 12:1–19 73. Chance B, Williams GR (1955) Respiratory enzymes in oxidative phosphorylation. J Biol 1:409–428 74. Chance B, Williams GR (1955) Respiratory enzymes in oxidative phosphorylation. IV. The respiratory chain. J Biol Chem 217: 429–438 75. Kuznetsov AV, Strobl D, Ruttmann E et al (2002) Evaluation of mitochondrial respiratory function in small biopsies of liver. Anal Biochem 305:186–194 76. Gnaiger E (2008) Polarographic oxygen sensors, the oxygraph, and high-resolution respirometry to assess mitochondrial function. In: Dykens JA, Will Y (eds) Drug-induced mitochondrial dysfunction. John Wiley & Sons, Hoboken, pp 327–351 77. Kuznetsov AV, Veksler V, Gellerich FN et al (2008) Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nat Protoc 3:965–976 78. Veksler VI, Kuznetsov AV, Sharov VG et al (1987) Mitochondrial respiratory parameters in cardiac tissue: a novel method of assessment by using saponin-skinned fibers. Biochim Biophys Acta 892:191–196 79. Saks VA, Veksler VI, Kuznetsov AV et al (1998) Permeabilized cell and skinned fiber techniques in studies of mitochondrial function in vivo. Mol Cell Biochem 184:81–100 80. Hutter E, Renner K, Pfister G et al (2004) Senescence-associated changes in respiration and oxidative phosphorylation in primary human fibroblasts. Biochem J 380:919–928

171

81. Keuper M, Jastroch M, Yi CX et al (2014) Spare mitochondrial respiratory capacity permits human adipocytes to maintain ATP homeostasis under hypoglycemic conditions. FASEB J 28:761–770 82. Divakaruni AS, Hsieh WY, Minarrieta L et al (2018) Etomoxir inhibits macrophage polarization by disrupting CoA homeostasis. Cell Metab 28:490–503.e7 83. Pesta D, Gnaiger E (2012) High-resolution respirometry: OXPHOS protocols for human cells and permeabilized fibers from small biopsies of human muscle. Methods Mol Biol 810:25–58 84. Brand MD, Chien LF, Diolez P (1994) Experimental discrimination between proton leak and redox slip during mitochondrial electron transport. Biochem J 297:27–29 85. Harper M, Brand D (1993) The quantitative contributions of mitochondrial proton leak and ATP turnover reactions to the changed respiration rates of hepatocytes from rats of different thyroid status. J Biol Chem 268: 14850–14860 86. Ruas JS, Siqueira-Santos ES, Amigo I et al (2016) Underestimation of the maximal capacity of the mitochondrial electron transport system in oligomycin-treated cells. PLoS One 11:1–20 87. Murphy MP (2001) How understanding the control of energy metabolism can help investigation of mitochondrial dysfunction, regulation and pharmacology. Biochim Biophys Acta 1504:1–11 88. Rolfe DFS, Brown GC (1997) Cellular energy utilization and molecular origin of standard metabolic rate in mammals. Physiol Rev. 77: 731–758 89. Jaber SM, Yadava N, Polster BM (2020) Mapping mitochondrial respiratory chain deficiencies by respirometry: beyond the mito stress test. Exp Neurol 328:113282 90. Larsen S, Nielsen J, Hansen CN et al (2012) Biomarkers of mitochondrial content in skeletal muscle of healthy young human subjects. J Physiol 590:3349–3360 91. Fell DA (1997) Understanding the control of metabolism. Ashgate Publishing, Surrey 92. Eigentler A, Draxl A, Wiethu¨chter A (2015) Laboratory protocol: citrate synthase a mitochondrial marker enzyme. Mitochondrial Physiol Netw 04:1–11 93. Salabei JK, Gibb AA, Hill BG (2014) Comprehensive measurement of respiratory activity in permeabilized cells using extracellular flux analysis. Nat Protoc 9:421–438

172

Matthew T. Lewis et al.

94. Kunz WS, Kuznetsov AV, Schulze W et al (1993) Functional characterization of mitochondrial oxidative phosphorylation in saponin-skinned human muscle fibers. Biochim Biophys Acta 1144:46–53 95. Kongas O, Yuen TL, Wagner MJ et al (2002) High Km of oxidative phosphorylation for ADP in skinned muscle fibers: where does it stem from? Am J Physiol Cell Physiol 283: 743–751 96. Gayeski TEJ, Honig CR (1986) O2 gradients from sarcolemma to cell interior in red muscle at maximal V˙O2. Am J Physiol 251(4 Pt 2): H789–H799 97. Metelkin E, Goryanin I, Demin O (2006) Mathematical modeling of mitochondrial adenine nucleotide translocase. Biophys J 90: 423–432 98. Saks VA, Kuznetsov A, Andrienko T et al (2003) Heterogeneity of ADP diffusion and regulation of respiration in cardiac cells. Biophys J 84:3436–3456 99. Wiseman RW, Jeneson JAL, Kushmerick MJ (1996) Why is the sensitivity of mitochondria to ADP over tenfold lower in permeabilized fibers than in vivo? Biothermokinetics living cell. Biothermokinetics Press, Amsterdam, pp 124–127 100. Saks VA, Belikova YO, Kuznetsov AV (1991) In vivo regulation of mitochondrial respiration in cardiomyocytes: specific restrictions for intracellular diffusion of ADP. Biochim Biophys Acta 1074:302–311 101. Scheibye-Knudsen M, Quistorff B (2009) Regulation of mitochondrial respiration by inorganic phosphate; comparing permeabilized muscle fibers and isolated mitochondria prepared from type-1 and type-2 rat skeletal muscle. Eur J Appl Physiol 105:279–287 102. Kuznetsov AV, Lassnig B, Margreiter R et al (1998) Diffusion limitation of oxygen versus ADP in permeabilized muscle fibers. In: Biothermokinetics post genomic era. Chalmers Reproservice, Go¨teborg, pp 273–276 103. Gnaiger E (2003) Oxygen conformance of cellular respiration: a perspective of mitochondrial physiology. Adv Exp Med Biol 543:39–55 104. Miotto PM, LeBlanc PJ, Holloway GP (2018) High-fat diet causes mitochondrial dysfunction as a result of impaired ADP sensitivity. Diabetes 67:2199–2205 105. Hart CR, Lanza IR (2018) Mitochondrial ADP sensitivity and transport: new insights into diet-induced mitochondrial impairments. Diabetes 67(11):2152–2153

106. Dirks ML, Miotto PM, Goossens GH et al (2020) Short-term bed rest-induced insulin resistance cannot be explained by increased mitochondrial H2O2 emission. J Physiol 598:123–137 107. Kasper JD (2018) The role and regulation of pyruvate dehydrogenase in skeletal muscle bioenergetics and type 2 diabetes. Diabetes Med J 38(3):181–186 108. Richardson RS, Noyszewski EA, Kendrick KF et al (1995) Myoglobin O2 desaturation during exercise: evidence of limited O2 transport. J Clin Invest 96:1916–1926 109. Rodenburg JB, De BRW, Jeneson JAL et al (1994) 31P-MRS and simultaneous quantification of dynamic human quadriceps exercise in a whole body MR scanner. J Appl Physiol 77:1021–1029 110. Whipp BJ, Rossiter HB, Ward SA et al (1999) Simultaneous determination of muscle and O2 uptake kinetics during whole body NMR spectroscopy. J Appl Physiol 86:742–747 111. Jaber Y, Jimenez Francisco E, Bartlett MF et al (2020) Magnetic resonance compatible knee extension ergometer. J Biomech Eng 142(9):095001 112. Ryschon TW, Fowler MD, Wysong RE et al (1997) Efficiency of human skeletal muscle in vivo: comparison of isometric, concentric, and eccentric muscle action. J Appl Physiol 83:867–874 113. Amara CE, Shankland EG, Jubrias SA et al (2007) Mild mitochondrial uncoupling impacts cellular aging in human muscles in vivo. Proc Natl Acad Sci U S A 104: 1057–1062 114. Amara CE, Marcinek DJ, Shankland EG et al (2008) Mitochondrial function in vivo: spectroscopy provides window on cellular energetics. Methods 46:312–318 115. McConnell DG (1965) The isolation of retinal outer segment fragments. J Cell Biol 27: 459–473 116. Greif RL, Alfano JA, Eich E (1966) Density gradient centrifugation of liver mitochondria from normal and thyrotoxic rats. Endocrinology 78:733–736 117. 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: 643–666 118. Messer JI, Jackman MR, Willis WT (2004) Pyruvate and citric acid cycle carbon requirements in isolated skeletal muscle mitochondria. Am J Physiol Cell Physiol 286(3): C565–C572

Chapter 11 Mitochondrial Toxicity of Organic Arsenicals Yu-Jiao Liu and Yi Liu Abstract Arsenic is either notorious toxicant or miracle cure for acute promyelocytic leukemia and several other diseases. It interacts with mitochondria directly or indirectly, by interacting with mitochondrial enzymes, such as respiratory chain complexes and tricarboxylic acid cycle proteins, or affecting mitochondrial homeostasis via ROS or mitochondrial outer membrane permeabilization. Given the ubiquitous presence of mitochondria and indispensable role in cellular metabolism, arsenical-mitochondrial interactions may manifest clinical importance by revealing mechanism of disease curation, preventing severe side effects, and foreseeing potential health issues. Here, we described the interaction between isolated mitochondria and arsenicals. Key words Isolated mitochondria, Organic arsenicals, Arsenic, ROS, Electron transport chain, Respiratory chain complex, Membrane permeability

1

Introduction Arsenic (As) is widely distributed in the environment. Soluble inorganic arsenite (As3+) and arsenate (As5+) are the predominant forms of arsenic contamination in groundwater. Inorganic arsenic (iAs) also presents in sulfide state within complex minerals with copper, iron, silver, etc. [1]. Classified as a Group I human carcinogen by the International Agency for Research on Cancer, Arsenic is released into the environment from geological, such as volcano eruptions and weathering of rocks and soils, and anthropogenic sources, like mining, agricultural irrigation and synthetic (organoarsenic) oAs usage as defoliants, herbicides, and antimicrobial growth promoters [2]. IAs is bio-transformed to various oAs species including arsenosugars, arsenobetaine, arsenolipids, monomethylarsonic acid (MMAIII/MMAV), and dimethylarsinic acid (DMAIII/DMAV), etc. by bacteria, fungi, and mammals including humans [3, 4]. In liver, glutathione (GSH)-mediated As5+ reduction and oxidative methylation of As3+ by arsenite methyltransferase (As3MT)

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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to pentavalent organic arsenic species (e.g., MMAV and DMAV) make up the metabolism of arsenic species. Under elevated concentrations of GSH in intracellular hepatocytes, the –OH group of As(OH)3 is sometimes replaced with glutathionyl moieties to form (GS)2AsOH [5]. Purine nucleoside phosphorylase (PNP) and glutathione S-transferase omega (GSTO; isoforms 1/2) catalyze arsenate (AsV) to arsenite (AsIII). S-adenosylmethionine (SAM) donates its methyl group to AsIII to complete methylation of AsIII. As3MT also involved in various methylated species of arsenic forming, such as monomethylarsonic acid (MMAIII/MMAV) and dimethylarsinic acid (DMAIII/DMAV). And their metabolism contributes to the toxicity of iAs even though methylation of iAs is thought to be a detoxification process, methylated arsenites are much more toxic than inorganic arsenic. It is widely accepted that the order of cytotoxicity was MMAIII > AsIII V V V
As > MMA ~ DMA . Long-lasting and/or high-dosage arsenic exposure damages the human cardiovascular, neurological, respiratory, dermal, hepatic, and reproductive systems [1, 3, 6, 7]. Despite being contaminant and toxicant, Arsenics have been recognized as medicinal compounds throughout human history [8–12]. Possessing high affinity for sulfhydryl groups especially vicinal dithiols [13], AsIII can conformationally and functionally affect numerous proteins, resulting dysfunction of protein folding, sulfur metabolism and redox balance [14, 15]. Arsenicals preferentially target the VDAC2 (voltage-dependent anion channel) on the mitochondrial membrane via covalent binding to the vicinal dithiols [13]. Mitochondrial respiration dysfunction occurs rapidly after As2O3 treatment with increased superoxide formation and decreased oxygen consumption. The preference for modification of cysteine also suggests that arsenite can affect the redox status within the cell. Two oAs induce the burst of ROS, thus consumption of antioxidants and high sensitivity for oxidants, which further induce the mitochondria dysfunction counting in the collapse of membrane potential, ATP level decline, membrane swelling, MPTP (mitochondrial permeability transition pore) opening, Ca2+ and cytochrome c release to trigger the mitochondria-dependent apoptosis [14]. To learn from the involvement of GSH in the metabolism of arsenic, GSAO (4-(N-(S-glutathionylacetyl)amino) phenylarsonous acid) was synthesized with phenylarsonous acid and GSH [16]. GSAO localized at mitochondria of cultured endothelial cells and binds two mitochondrial matrix cysteines of ANT, triggering MPTP opening in isolated mitochondria followed by apoptosis cascade reactions [17, 18]. To further consider the methylation of arsenic, S-dimethylarsinoglutathione (darinaparsins, ZIO-101) was the structural optimization of GSAO with dimethylation of As of GSAO [17]. Darinaparsin was shown to produce a

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higher amount of reactive oxygen than As2O3. And both darinaparsin and As2O3 induced upregulation of BH3-only proteins to promote mitochondrial-dependent apoptosis [19]. Given the tight relationship of arsenicals and mitochondria, mito-targeted arsenicals were synthesized. Even short-time incubation of PDT-PAO-TPP can disturb respiration by inhibition of respiratory chain complex I and IV, causing over death threshold ROS level to initiate apoptosis [20]. Besides, fluorescent PDT-PAO-F16 accumulated into mitochondria within 1 h and suppressed the activity of pyruvate dehydrogenase complex (PDHC) leading to ATP synthesis paralysis and thermogenesis disorder [21]. Even the non-mito-targeted arsenicals interact with mitochondria either in the upstream or the downstream of their mechanism pathways. The lactate dehydrogenase A (LDHA) inhibitory arsenical converted the glycolysis to oxidative phosphorylation causing ROS burst and mitochondrial dysfunction [22]. A series of arsenicals interact with mitochondria through ROS or intrinsic apoptosis with collapsed mitochondrial membrane potential, reduced ATP level, cytochrome c release, etc. [15, 23]. To decipher the deleterious or anti-cancer effects of organic arsenicals, it is necessary to investigate not only their cellular function but also into the individual organelles, which are themselves the autonomous regulatory bodies functioning in an orchestrated manner. Mitochondria are essential for life. As the most important organelle, on the one hand, mitochondria govern energy transformation and ATP production through the tricarboxylic acid cycle (TCA) and oxidative phosphorylation (OXPHOS) for cell growth and proliferation [24, 25]. On the other hand, mitochondria, regulating the signal transmission [26, 27], are paradoxically essential for cell death, such as apoptosis, necroptosis, ferroptosis and pyroptosis [28] and have been associated with numerous diseases, such as Leigh syndrome, Huntington’s disease, Alzheimer’s disease [26, 29–33]. Mitochondria are constituted of mitochondrial matrix, inner mitochondrial membrane (IMM, or cristae), intermembrane space (IMS), and outer mitochondrial membrane (OMM). IMM are intensively folded and well-controlled structure where complexes of the respiratory chain (RC) (complexes I–IV) are located. OMM is responded for the communication with cytosol through porous structures, like the voltage-dependent anion channel (VDAC) [34]. About 85% of cytochrome c and 94% of complex III and ATP synthase are stored in cristae. Mitochondrial well-defined but plastic structure is coupled to their various functions when needed [35]. By fusion and fission, mitochondria can switch from an elongated and interconnected states to fragmented states [36]. Although there is a plenty of

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room to discover how cristae shapes, ROS, the NADH/NAD ratio, oxygen concentration, and the ATP/ADP ratio can be important signaling molecules that crosstalk between mitochondrial metabolism, cellular pathways, and mitochondrial cristae remodeling. Mitochondrial fission was related to mitochondrial apoptosis and was also implied to be a prerequisite for mitophagy, while mitochondria fusion contributed to increased mitochondrial metabolism [37]. Mitophagy control mitochondrial quality by degrading dysfunctional mitochondria with sequestration them into autophagosomes [38]. When cells encounter nutrient deprivation, mitophagy is activated to prevent mitochondrial degradation and to promote ATP production by mitochondrial fusion. A group of GTP-dependent dynamin-like proteins involved in the fusion and fission, such as Mitofusin 1,2, the pro-fusion proteins Optic atrophy 1 (OPA1), the pro-fission Drp1, and structural proteins, such as the Prohibitin (PHB) family proteins and the mitochondrial contact site and cristae-organizing system (MICOS) complex [39]. The tricarboxylic acid (TCA) cycle consists of eight enzymatic reactions where electrons are transferred to nicotinamide adenine dinucleotide (NAD) and flavin adenine dinucleotide (FAD) generating reduced NADH and FADH2. NADH and FADH2 are then passed to RC for oxidative phosphorylation (OXPHOS). Complex I (CI) or NADH-coenzyme Q oxidoreductase is a large protein complex built of 46 subunits, and NADH dehydrogenase (ND) subunits 1–7 (ND1–ND7) are mitochondrially encoded [40]. Electrons were transferred from matrix NADH to ubiquinone (CoQ) by CI generating NAD and ubiquinol (QH2), respectively. Complex II (CII) or succinate dehydrogenase (SDH) is built of four subunits (SDHA-D), all encoded by nuclear genes [41]. CII catalyzes the oxidation of succinate to fumarate by giving electron to FAD forming FADH2, followed by oxidation of FADH2 and transferring electrons to CoQ generating QH2 [42]. Complex III (CIII) or CoQ-cytochrome c reductase is a cytochrome bc1 complex which transfers the electrons to cytochrome c from QH2. CIII is a symmetrical dimer with 11 subunits per monomer. A high-potential (2Fe-2S) cluster wrapped by an ironsulfur protein, cytochrome b (bL and bH), and cytochrome c1 make up the catalytically active subunits [43]. CIII transfer electrons by the Q-cycle to cytochrome c. Complex IV (CIV) or cytochrome c oxidase is responsible for electron transfer from cytochrome c to the terminal electron acceptor O2 generating H2O. Mammalian CIV consists of 13 different subunits with 3 core subunits (I, II, and III) encoded by mitochondrial DNA and others encoded by nuclear genes [44]. The non-core subunits modulate CIV activity according to the ATP/ADP-ratio. Electrons are transferred from cytochrome c to subunit II, heme, subunit I and finally O2 generating H2O.

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Complex V (CV) or F1F0 ATP synthase consists of F0 domain and F1 domain located in the IMM and mitochondrial matrix, respectively. When 2 electrons are transferred from CI to CIII, and then CIV, 4, 4, and 2 protons are removed from the matrix through CI, CIII, and CIV, respectively or 0, 4, 2 protons through CII, CIII,and CIV, respectively [40]. Protons from IMS cross CV leading to F1F0 ATP synthase conformational changes where ADP is phosphorylated to form ATP. The proton gradient across the IMM highly negatively polarize IMM and is determined by the mitochondrial membrane potential (MMP, ΔΨm). Due to the downregulated CV in cancer mitochondria causing accumulation of protons in IMS, MMPs in cancer cells (~220 mV) are about 60 mV higher than those in normal cells (108–180 mV) [45]. As 85–90% of intracellular O2 is consumed for ATP production in normal cells, mitochondria maintain low oxygen levels (0.5–200 μM). Owing to the hypoxic tumoral environment, tumor cells have lower concentration of intracellular O2. However, decreased ATP synthesis in cancer cells attribute 2.5-fold lower oxygen consumption rate (OCR) than that in normal cells, leading to higher mitochondrial oxygen level in cancer cells [46]. Besides, ROS are produced during normal cellular metabolism mainly by Complex I and III where about 2% of the electrons are leaked to react with O2 generating superoxide. Owing to more leaked electrons under hypoxic condition and surplus O2, cancer cell possess ~0.1 mM ROS while normal cells only have ~20 nM [47, 48]. Although normal level ROS are essential for maintaining proper cellular signaling, burst ROS, toxic to lipids, proteins and nucleic acids, may cause cellular dysfunction and associate with a wide range of diseases. And about 80% O2•generated by complex I and III is released into the IMS and others into mitochondrial matrix. Electron transport chain in mitochondria is the main resource of intracellular ROS, mitochondria are also the target of ROS. When ROS are elevated above the threshold of cell death, mitochondrial dysfunction occurs and successive depolarization of the MMP leading to releasing cytochrome c to initiate apoptosis. Therefore, monitoring mitochondrial oxygen consumption rate and ROS level are essential to estimate cell status under certain condition. When calcium overload in the mitochondrial matrix, or cysteine residues of adenine-nucleotide translocase (ANT) in the IMM undergo alkylation and/or oxidation, ANT involves in the formation of a non-specific channel named mitochondrial permeability transition pore (MPTP) allowing all solutes of up to 1.5 kDa permeabilization from both sides. MPTP consists of ANT, VDAC, cyclophilin D, and others (Bcl-2 family proteins, mitochondrial hexokinase, and mitochondrial creatine kinase) [49].

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BAX and, to a lesser degree, BAK shuttle between the mitochondria and cytoplasm under normal condition. During apoptosis, BAX and BAK can be directly activated by binding BH3-only proteins called direct activators (BID, PUMA, and BIM) resulting in their stabilization at OMM and their homodimerization. BAX/BAK dimers further oligomerize thus forming higher-order multimers that generate lipid pores within the OMM causing mitochondrial outer membrane permeabilization (MOMP) [28]. The MPTP could allow the release of soluble IMS proteins, for instance, cytochrome c, to cytosol, which can be further facilitated by IMM remodeling or cristae remodeling involving releasing of cytochrome c from mitochondrial cristae. The pore forming by BAX/BAK could enable IMM extrusion through the OMM, whereupon the IMM herniates and ruptures leading to releasing of mitochondrial DNA (mtDNA) [28, 39]. Given the importance of mitochondria and the tight relationship between arsenicals and mitochondria, herein, we isolated mitochondria from mice liver and investigated the interaction between mitochondria and arsenicals.

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Methods 1. Mitochondria isolation. Fresh liver from Wistar rats in precooled beaker was minced quickly and washed with solution A (0.07 M sucrose, 0.22 M mannitol, 0.02 M HEPES, 2 mM Tris-HCl, 1 mM EDTA-2Na, 0.4% BSA, pH 7.4) three times. The remainder was suspended in solution A (20 mL per gram liver) and homogenized in a precooled Dounce Tissue Grinder (WHEATON). The homogenate was centrifuged at 3000  g for 2 min followed by supernatant centrifugation at 17,500  g for 5 min. Then the pellet was washed by solution A and then solution B (0.07 M sucrose, 0.22 M mannitol, 0.01 M TrisHCl, 1 mM EDTA, pH 7.4) at 17,500  g for 5 min before being suspended in solution B0 (0.22 M mannitol, 0.07 M sucrose, 1 mM EDTA, pH 7.4, 0.6 mL per gram liver). The mitochondrial protein concentration was determined by the Biuret method with serum albumin as standard. All the operations above were performed aseptically at 0–4  C and the mitochondria were of good quality with 3 h. 2. Mitochondrial swelling. Mitochondria (0.5–0.8 mg protein per mL) were suspended in 2 mL or 200 μL of MAB solution (0.2 M sucrose, 10 mM Tris-HCl, 10 mM MOPS, 10 μM EGTA, 1 mM Na3PO4, 3 μg/mL Oligomycin, 5 mM succinate, and 2 μM rotenone, pH 7.2). Agents of interest at related concentrations

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or related solvent of agents were added for the following tests. The absorbance at 540 nm for 8 or 60 min at 25  C was monitored by ultraviolet-visible spectrophotometer (UNICO, USA) or multimode plate reader VICTOR™ X5 (PE, USA). 3. Mitochondrial membrane potential (ΔΨm). Mitochondria (0.5–0.8 mg protein per mL) were suspended in 2 mL or 200 μL MAB solution adding 250 μM Rh123, with/without agents of interest. The fluorescence emission intensity of Rh123 was monitored by multimode plate reader VICTOR™ X5 (PE, USA). 4. H+ and K+ permeabilization. Mitochondria (0.5–0.8 mg protein per mL) were suspended in 2 mL or 200 μL of solution H (135 mM potassium acetate, 5 mM HEPES, 0.1 mM EGTA, 0.2 mM EDTA, 1 μg/ mL valinomycin, and 2 μM rotenone, pH ¼ 7.1) for permeabilization of H+ or solution K (135 mM KNO3, 5 mM HEPES, 0.1 mM EGTA, 0.2 mM EDTA, and 2 μM rotenone, pH ¼ 7.1) for permeabilization of K+. Then agents of interest were added to the mitochondrial suspension before detection of absorbance at 540 nm for 8 min at 25  C by ultravioletvisible spectrophotometer (UNICO, USA) or multimode plate reader VICTOR™ X5 (PE, USA). Valinomycin was added into solution H to ensure that K+ had passed completely through the mitochondrial inner membrane. 5. Membrane fluidity. Mitochondria (0.5–0.8 mg protein per mL) were suspended in 2 mL MAB solution adding 2 μM hematoporphyrin (HP), with/without agents of interest. The anisotropy value was recorded by LS 55 fluorescence spectrophotometer (PE, USA) under magnetic stirring at 25  C (λex ¼ 520 nm, λem ¼ 626 nm). 6. Respiration. Mitochondria (1 mg/mL) suspended in MRB solution (100 mM sucrose, 10 mM Tris-HCl, 10 mM MOPS, 2 mM MgCl2, 50 mM KCl, 10 mM K2HPO4, 1 mM EDTA, and 2 μM rotenone, pH 7.4) were incubated with agents of interest for certain time or were mixed with agents immediately before tests. Oxygen consumption rate (OCR) was recorded by Clark Oxygen Electrode (Hansatech Instruments, Norfolk, UK) for 1–2 min before the addition of 1 M succinate to initiate State 4. One to two minutes later state 3 was initiated by adding 1 M succinate and 250 μM ADP into the above solution followed by initiation of uncoupled respiration by the addition of 3 mM DNP. The OCR was recorded under magnetic stirring at 25  C without stop during the addition of agents to initiate the three

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states. Respiration control ratio was the ratio of OCR of state 3 and that of state 4. To note: try the best to complete the experiments in the same time and get data from the same time periods. 7. Microcalorimetry. One milliliter solution B0 containing mitochondria (5 mg/ mL), 15 mM pyruvate, with/without agents of interest were added into an ampoule at 4  C under sterile condition. The ampoule was sealed and warmed in hands before loading into TAM III (TAM III, TA Instruments, New Castle, USA). The testing temperature was set at 30  C. When the samples were heated to 30  C, thermogenic curves were recorded and terminated when the signal went back to ground. To note: add all the things into ampoules under sterile condition. Try the best to warm the sample to 30  C as soon as possible and keep the ampoules away from any contaminants even sweat or grease in hands or gloves. 8. Evaluation of respiratory chain complex I, II, III, and IV activities. Mitochondria (16–20 μg/mL) in 200 μL related assay buffer were incubated with/without agents of interest or inhibitor for 3–5 min before tests which were monitored by multimode microplate reader (Tecan Spark® 10 M, Switzerland) for at least 3 min. Specific inhibitors, namely CI (rotenone, 5 μM), CII (sodium malonate, 10 mM), CIII (antimycin, 4 μM), and CIV (NaN3, 3 mM), were tested to determine the specificity of the respiratory complexes activities. CI: mitochondria were incubated firstly with agents of interest for 3 min in 10 mM Tris-HCl buffer (pH 8.0), followed by addition of 80 μM 2,3-dimethoxy-5-methyl-6-decyl1,4-benzoquinone (DB), 1 mg/mL BSA, 3 mM NaN3, and 0.4 μM antimycin for another 5 min. Then NADH (200 μM) was added to initiate NADH oxidation by CI which was recorded by absorption at 340 nm immediately. CII: mitochondria were incubated firstly with agents of interest for 3 min in 50 mM PBS (pH 7.4), followed by addition of 10 mM succinate, 50 μM 2,6-dichlorophenolindophenol (DCPIP), 3 mM NaN3, 2 μM rotenone, and 2 μM antimycin. Then 20 μM DB was added to initiate reduction of DCPIP in association with CII-catalyzed DB reduction recorded by absorbance at 600 nm. CIII: mitochondria were incubated with agents of interest for 3 min in 50 mM Tris-HCl buffer (pH 7.4) with 1 mM EDTA, 250 mM sucrose, 3 mM NaN3, and 30 μM oxidized cytochrome c. Then 80 μM ubiquinol was added and the reduction of cytochrome c was measured according to absorbance at 550 nm.

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CIV: mitochondria were incubated with agents of interest for 3 min in 10 mM Tris-HCl (pH 7.0) with 25 mM sucrose, 120 mM KCl, and 0.025% n-dodecyl-h-D-maltoside. After addition of 50 μM reduced cytochrome c, the oxidation of cytochrome c was measured by absorbance change at 550 nm. To note: conduct all the experiments as quick as possible and keep all the materials sterile. 9. Transmission electron microscopy of mitochondria. Mitochondria (0.5 mg/mL) incubated with agents of interest for certain time (usually less than 1 h) were fixed by 2.5% glutaraldehyde solution before preparation for transmission electron microscope (Tecnai G20 Twin, FEI, Hillsboro, USA). Mitochondria extracted from rat liver maintained the integrity of the classical ultrastructure: compact cristae, narrow IMS, and well-defined outer membrane. After mitochondria interacted with organic arsenicals, mitochondrial underwent swelling with declined electron density and even empty membrane vesicles, membranes broken with irregular and even ruptured IMM and OMM. These phenomena can be coherent with results of swelling, H+ and K+ permeabilization and even membrane potential. To note, the mitophagy, fission or fusion only can be seen in cells as they are cascade reactions in cells. 10. Measurement of lipid peroxidation. Mitochondrial membrane lipid peroxidation was assessed by the consumption of oxygen using a Clark Oxygen Electrode. Mitochondria were injected into 1 mL lipid peroxidation medium (agents of interest, 175 mM KCl, 10 mM TrisHCl, and 3 μM rotenone, pH 7.4). Membrane lipid peroxidation was initiated by the addition of 1 mM ADP/0.1 mM Fe2+. To note: The iron(II) solution must be prepared before using.

Acknowledgments We gratefully acknowledge the financial support of National Natural Science Foundation of China (22073070, 21673166). References 1. Sharma VK, Sohn M (2009) Aquatic arsenic: toxicity, speciation, transformations, and remediation. Environ Int 35(4):743–759. https:// doi.org/10.1016/j.envint.2009.01.005 2. Tam LM, Price NE, Wang Y (2020) Molecular mechanisms of arsenic-induced disruption of

DNA repair. Chem Res Toxicol 33(3): 709–726. https://doi.org/10.1021/acs. chemrestox.9b00464 3. Mandal BK, Suzuki KT (2002) Arsenic round the world: a review. Talanta 58(1):201–235.

182

Yu-Jiao Liu and Yi Liu

https://doi.org/10.1016/S0039-9140(02) 00268-0 4. Taylor V, Goodale B, Raab A, Schwerdtle T, Reimer K, Conklin S, Karagas MR, Francesconi KA (2017) Human exposure to organic arsenic species from seafood. Sci Total Environ 580: 266–282. https://doi.org/10.1016/j. scitotenv.2016.12.113 5. Ali W, Zhang H, Junaid M, Mao K, Xu N, Chang C, Rasool A, Wajahat Aslam M, Ali J, Yang Z (2020) Insights into the mechanisms of arsenic-selenium interactions and the associated toxicity in plants, animals, and humans: a critical review. Crit Rev Environ Sci Technol 51(7):704–750. https://doi.org/10.1080/ 10643389.2020.1740042 6. Druwe IL, Vaillancourt RR (2010) Influence of arsenate and arsenite on signal transduction pathways: an update. Arch Toxicol 84(8): 585–596. https://doi.org/10.1007/s00204010-0554-4 7. Abernathy CO, Liu YP, Longfellow D, Aposhian HV, Beck B, Fowler B, Goyer R, Menzer R, Rossman T, Thompson C, Waalkes M (1999) Arsenic: health effects, mechanisms of actions, and research issues. Environ Health Perspect 107(7):593–597. https://doi.org/ 10.2307/3434403 8. Kumagai Y, Sumi D (2007) Arsenic: signal transduction, transcription factor, and biotransformation involved in cellular response and toxicity. Annu Rev Pharmacol Toxicol 47(1):243–262. https://doi.org/10.1146/ annurev.pharmtox.47.120505.105144 9. Zhu J, Chen Z, Lallemand-Breitenbach V, de The H (2002) How acute promyelocytic leukaemia revived arsenic. Nat Rev Cancer 2(9): 705–714. https://doi.org/10.1038/nrc887 10. Diaz Z, Mann KK, Marcoux S, Kourelis M, Colombo M, Komarnitsky PB, Miller WH Jr (2008) A novel arsenical has antitumor activity toward As2O3-resistant and MRP1//ABCC1overexpressing cell lines. Leukemia 22(10): 1853–1863. https://doi.org/10.1038/leu. 2008.328 11. Noguera NI, Piredda ML, Taulli R, Catalano G, Angelini G, Gaur G, Nervi C, Voso MT, Lunardi A, Pandolfi PP, Lo-Coco F (2016) PML/RARa inhibits PTEN expression in hematopoietic cells by competing with PU.1 transcriptional activity. Oncotarget 7(41): 66386–66397. https://doi.org/10.18632/ oncotarget.11964 12. Gibaud S, Jaouen G (2010) Arsenic-based drugs: from fowler’s solution to modern anticancer chemotherapy. In: Jaouen G, MetzlerNolte N (eds) Medicinal organometallic

chemistry. Springer, Berlin, pp 1–20. https:// doi.org/10.1007/978-3-642-13185-1_1 13. Yang Z, Kang DH, Lee H, Shin J, Yan W, Rathore B, Kim H-R, Kim SJ, Singh H, Liu L, Qu J, Kang C, Kim JS (2018) A fluorescent probe for stimulated emission depletion super-resolution imaging of vicinal-dithiolproteins on mitochondrial membrane. Bioconjug Chem 29(4):1446–1453. https://doi.org/ 10.1021/acs.bioconjchem.8b00128 14. Fan XY, Liu YJ, Chen K, Jiang FL, Hu YJ, Liu D, Liu Y, Ge YS (2018) Organic arsenicals target thioredoxin reductase followed by oxidative stress and mitochondrial dysfunction resulting in apoptosis. Eur J Med Chem 143: 1090–1102. https://doi.org/10.1016/j. ejmech.2017.05.022 15. Liu Y, Duan D, Yao J, Zhang B, Peng S, Ma H, Song Y, Fang J (2014) Dithiaarsanes induce oxidative stress-mediated apoptosis in HL-60 cells by selectively targeting thioredoxin reductase. J Med Chem 57(12):5203–5211. https://doi.org/10.1021/jm500221p 16. Elliott MA, Ford SJ, Prasad E, Dick LJ, Farmer H, Hogg PJ, Halbert GW (2012) Pharmaceutical development of the novel arsenical based cancer therapeutic GSAO for phase I clinical trial. Int J Pharm 426(1):67–75. https://doi.org/10.1016/j.ijpharm.2012. 01.024 17. Park D, Chiu J, Perrone GG, Dilda PJ, Hogg PJ (2012) The tumour metabolism inhibitors GSAO and PENAO react with cysteines 57 and 257 of mitochondrial adenine nucleotide translocase. Cancer Cell Int 12(1):11–16. https:// doi.org/10.1186/1475-2867-12-11 18. Don AS, Kisker O, Dilda P, Donoghue N, Zhao X, Decollogne S, Creighton B, Flynn E, Folkman J, Hogg PJ (2003) A peptide trivalent arsenical inhibits tumor angiogenesis by perturbing mitochondrial function in angiogenic endothelial cells. Cancer Cell 3(5):497–509. https://doi.org/10.1016/S1535-6108(03) 00109-0 19. Matulis SM, Morales AA, Yehiayan L, Croutch C, Gutman D, Cai Y, Lee KP, Boise LH (2009) Darinaparsin induces a unique cellular response and is active in an arsenic trioxide-resistant myeloma cell line. Mol Cancer Ther 8(5):1197–1206. https://doi.org/ 10.1158/1535-7163.MCT-08-1072 20. Fan XY, Liu YJ, Cai YM, Wang AD, Xia YZ, Hu YJ, Jiang FL, Liu Y (2019) A mitochondriatargeted organic arsenical accelerates mitochondrial metabolic disorder and function injury. Bioorg Med Chem 27(5):760–768. https://doi.org/10.1016/j.bmc.2019.01.008

Mitochondrial Toxicity of Organic Arsenicals 21. Liu YJ, Fan XY, Zhang DD, Xia YZ, Hu YJ, Jiang FL, Zhou FL, Liu Y (2020) Dual inhibition of pyruvate dehydrogenase complex and respiratory chain complex induces apoptosis by a mitochondria-targeted fluorescent organic arsenical in vitro and in vivo. ChemMedChem 15(6):552–558. https://doi.org/10.1002/ cmdc.201900686 22. Liu YJ, Fan XY, Wang AD, Xia YZ, Fu WR, Liu JY, Jiang FL, Liu Y (2019) LDHA suppression altering metabolism inhibits tumor progress by an organic arsenical. Int J Mol Sci 20(24): 6239–6256. https://doi.org/10.3390/ ijms20246239 23. Zhang D, Liu Y, Luo Z, Chen Y, Xu A, Liang Y, Wu B, Tong X, Liu X, Shen H, Liu L, Wei Y, Zhou H, Liu Y, Zhou F (2019) The novel thioredoxin reductase inhibitor A-Z2 triggers intrinsic apoptosis and shows efficacy in the treatment of acute myeloid leukemia. Free Radic Biol Med 146:275–286. https://doi.org/10.1016/j.freeradbiomed. 2019.11.013 24. Frazier AE, Thorburn DR, Compton AG (2019) Mitochondrial energy generation disorders: genes, mechanisms, and clues to pathology. J Biol Chem 294(14):5386–5395. https://doi.org/10.1074/jbc.R117.809194 25. Wallace DC, Fan W, Procaccio V (2010) Mitochondrial energetics and therapeutics. Annu Rev Pathol Mech Dis 5(1):297–348. https:// doi.org/10.1146/annurev.pathol.4.110807. 092314 26. Eisenberg L, Eisenberg-Bord M, EisenbergLerner A, Sagi-Eisenberg R (2020) Metabolic alterations in the tumor microenvironment and their role in oncogenesis. Cancer Lett 484:65– 71. https://doi.org/10.1016/j.canlet.2020. 04.016 27. Chi X, Wang Y, Heng BC, Wei Y, Guo Y, Zhang X, Zhao H, Yin Y, Deng X (2020) Mitochondria transfer enhances proliferation, migration, and osteogenic differentiation of bone marrow mesenchymal stem cell and promotes bone defect healing. Stem Cell Res Ther 11(1):245–260. https://doi.org/10.1186/ s13287-020-01704-9 28. Bock FJ, Tait SWG (2020) Mitochondria as multifaceted regulators of cell death. Nat Rev Mol Cell Biol 21(2):85–100. https://doi.org/ 10.1038/s41580-019-0173-8 29. Weidling IW, Swerdlow RH (2020) Mitochondria in Alzheimer’s disease and their potential role in Alzheimer’s proteostasis. Exp Neurol 330:113321–113328. https://doi.org/10. 1016/j.expneurol.2020.113321 30. Martı´nez-Reyes I, Cardona LR, Kong H, Vasan K, McElroy GS, Werner M, Kihshen H,

183

Reczek CR, Weinberg SE, Gao P, Steinert EM, Piseaux R, Budinger GRS, Chandel NS (2020) Mitochondrial ubiquinol oxidation is necessary for tumour growth. Nature 585:288–292. https://doi.org/10.1038/s41586-0202475-6 31. Jovanovski A, Pergolizzi B, Pironi L, Stanga S, Panuzzo C, Fava C, Cilloni D (2020) Mitochondria: a galaxy in the hematopoietic and leukemic stem cell universe. Int J Mol Sci 21(11):3928–3954. https://doi.org/10. 3390/ijms21113928 32. Jimenez-Blasco D, Busquets-Garcia A, Hebert-Chatelain E, Serrat R, VicenteGutierrez C, Ioannidou C, Go´mez-Sotres P, Lopez-Fabuel I, Resch-Beusher M, Resel E, Arnouil D, Saraswat D, Varilh M, Cannich A, Julio-Kalajzic F, Bonilla-Del Rı´o I, Almeida A, Puente N, Achicallende S, Lopez-Rodriguez M-L, Jolle´ C, De´glon N, Pellerin L, Josephine C, Bonvento G, Panatier A, Lutz B, Piazza P-V, Guzmán M, Bellocchio L, Bouzier˜ os JP, Marsicano G Sore A-K, Grandes P, Bolan (2020) Glucose metabolism links astroglial mitochondria to cannabinoid effects. Nature 583:603–608. https://doi.org/10.1038/ s41586-020-2470-y 33. Picard M, Wallace DC, Burelle Y (2016) The rise of mitochondria in medicine. Mitochondrion 30:105–116. https://doi.org/10. 1016/j.mito.2016.07.003 34. Vuda M, Kamath A (2016) Drug induced mitochondrial dysfunction: mechanisms and adverse clinical consequences. Mitochondrion 31:63–74. https://doi.org/10.1016/j.mito. 2016.10.005 35. Feng Q, Kornmann B (2018) Mechanical forces on cellular organelles. J Cell Sci 131(21):218479–218487. https://doi.org/ 10.1242/jcs.218479 36. Zhang L, Trushin S, Christensen TA, Bachmeier BV, Gateno B, Schroeder A, Yao J, Itoh K, Sesaki H, Poon WW, Gylys KH, Patterson ER, Parisi JE, Diaz Brinton R, Salisbury JL, Trushina E (2016) Altered brain energetics induces mitochondrial fission arrest in Alzheimer’s disease. Sci Rep 6(1):18725–18736. https://doi. org/10.1038/srep18725 37. Chen G, Li W, Kepp O, Zhu Y, Ma K, Chen Q (2020) Mitophagy, mitochondrial homeostasis, and cell fate. Front Cell Dev Biol 8:467– 480. https://doi.org/10.3389/fcell.2020. 00467 38. Han S, Jeong YY, Sheshadri P, Su X, Cai Q (2020) Mitophagy regulates integrity of mitochondria at synapses and is critical for synaptic maintenance. EMBO Rep 21(9):e49801. https://doi.org/10.15252/embr.201949801

184

Yu-Jiao Liu and Yi Liu

39. Cogliati S, Enriquez JA, Scorrano L (2016) Mitochondrial cristae: where beauty meets functionality. Trends Biochem Sci 41(3): 261–273. https://doi.org/10.1016/j.tibs. 2016.01.001 40. Ru-Zhou Z, Jiang S, Zhang L, Zhi-Bin Y (2019) Mitochondrial electron transport chain, ROS generation and uncoupling (Review). Int J Mol Med 44(2):3–15 41. Dalla Pozza E, Dando I, Pacchiana R, Liboi E, Scupoli MT, Donadelli M, Palmieri M (2020) Regulation of succinate dehydrogenase and role of succinate in cancer. Semin Cell Dev Biol 98:4–14. https://doi.org/10.1016/j. semcdb.2019.04.013 42. Pfanner N, Warscheid B, Wiedemann N (2019) Mitochondrial proteins: from biogenesis to functional networks. Nat Rev Mol Cell Biol 20(5):267–284. https://doi.org/10.1038/ s41580-018-0092-0 43. Schgger H, Link TA, Engel WD, Von JG (1986) Isolation of the eleven protein subunits of the bc1 complex from beef heart. Methods Enzymol 126(126):224–237. https://doi. org/10.1016/S0076-6879(86)26024-3 44. Kadenbach B, Hu¨ttemann M (2015) The subunit composition and function of mammalian cytochrome c oxidase. Mitochondrion 24(1):

64–76. https://doi.org/10.1016/j.mito. 2015.07.002 45. Forrest MD (2015) Why cancer cells have a more hyperpolarised mitochondrial membrane potential and emergent prospects for therapy. bioRxiv:025197. https://doi.org/10.1101/ 025197 46. Kueh Hao Y, Niethammer P, Mitchison Timothy J (2013) Maintenance of mitochondrial oxygen homeostasis by cosubstrate compensation. Biophys J 104(6):1338–1348. https:// doi.org/10.1016/j.bpj.2013.01.030 47. Waypa GB, Marks JD, Guzy R, Mungai PT, Schriewer J, Dokic D, Schumacker PT (2010) Hypoxia triggers subcellular compartmental redox signaling in vascular smooth muscle cells. Circ Res 106(3):526–535. https://doi. org/10.1161/CIRCRESAHA.109.206334 48. Cho H, Cho Y-Y, Shim MS, Lee JY, Lee HS, Kang HC (2020) Mitochondria-targeted drug delivery in cancers. Biochim Biophys Acta Mol Basis Dis 1866(8):165808–165827. https:// doi.org/10.1016/j.bbadis.2020.165808 49. Don AS, Hogg PJ (2004) Mitochondria as cancer drug targets. Trends Mol Med 10(8): 372–378. https://doi.org/10.1016/j. molmed.2004.06.005

Chapter 12 Measurements of Mitochondrial Respiration in Intact Cells, Permeabilized Cells, and Isolated Tissue Mitochondria Using the Seahorse XF Analyzer Jessica Pfleger Abstract Energy homeostasis is critical for cellular function. Significant increases in energy demand or reduced energy supply, however, often result in cellular dysfunction and death. Since mitochondria are the primary cellular energy source, their impairment is often pathogenic. Accordingly, quantitative measurements of cellular and mitochondrial energy utilization and production are crucial for understanding disease development and progression. In the final step of cellular respiration, specifically, oxidative phosphorylation within the mitochondria, oxygen is consumed and drives ATP production. Herein, we provide the complete protocols for measuring oxygen consumption rates and their coupling to ATP production in intact and permeabilized cells, as well as in mitochondria isolated from tissue using the Seahorse XF Extracellular Flux Analyzer (Agilent Technologies). Key words Cellular respiration, Mitochondrial respiration, Permeabilized cells, Bioenergetics, Seahorse XF Extracellular Flux Analyzer

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Introduction Optimally functioning mitochondria are crucial to the health of most cells. As the primary source of cellular energy, mitochondria are able to sense stress signals and subsequently enhance or reduce energy production, as well as initiate various cellular survival or death pathways [1]. Thus, mitochondrial heath is a direct indicator of cellular health. There are multiple ways to assess mitochondria quality, including examination of mitochondrial number, morphology, dynamics, membrane potential, ion handling, membrane integrity, and metabolism [2]. All of these processes, however, converge upon bioenergetics or the utilization and production of energy by the mitochondria. Accordingly, bioenergetic measurements are perhaps the most direct indicators of mitochondrial function and health.

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Assessment of cellular or mitochondrial bioenergetics include measurements of the endpoints of respiration, namely oxygen consumption and ATP production. Traditionally, the oxygen or Clark electrode is used to measure oxygen consumption rates in cells or mitochondria under various conditions [3], while ATP content is often measured using phosphorus-31 nuclear magnetic resonance spectroscopy or bioluminescence-based assays [2]. Herein, we will focus on measurements of oxygen consumption rates and the extrapolation of ATP production from these measurements. One major shortcoming of the oxygen electrode is that it requires cellular or mitochondrial suspensions for quantification [3], which is not ideal for adherent cells whose optimal function and survival depends on adhesion to extracellular matrix proteins and cell–cell interactions. Furthermore, dissociation of cells or isolation of mitochondria can cause damage or selection bias, particularly following experimental versus control treatment. A newer technology, the Seahorse XF Extracellular Flux Analyzer (Agilent Technologies), has overcome these limitations by allowing for measurements of oxygen consumption in monolayers of adherent cells [4]. In fact, these instruments, which are available in 8-, 24-, and 96-well formats, are able to simultaneously measure oxygen and pH levels in the assay medium in real time, allowing for the calculation of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR), respectively. In addition, multiple tests can be performed via injection of various metabolic substrates or compounds into the medium using the instrument’s four injection ports [4]. In this chapter, we outline two assays, the Mitochondrial Stress Test (Fig. 1) and the Mitochondrial Coupling Assay (Fig. 2), which assess similar bioenergetic parameters in intact cells and permeabilized cells or isolated mitochondria, respectively, as previously described [4–8]. Briefly, in the Mitochondrial Stress Test, baseline OCR is recorded and plotted versus time. Next, oligomycin, an inhibitor of the ATP synthase proton channel (FO subunit), is injected revealing the amount of the baseline OCR that contributes to ATP production (ATP-linked respiration), as well as the amount that is the result of proton leak. Trifluoromethoxy carbonyl cyanide phenylhydrazone (FCCP), a proton ionophore, is then used to dissipate the mitochondrial electrochemical gradient and uncouple oxidative phosphorylation, uncovering the maximal respiratory capacity (MRC) or maximal OCR of the cells. The MRC can also be used to calculate the spare or reserve respiratory capacity (RRC), which is the MRC minus the basal respiration. Finally, a combination of antimycin A and rotenone, electron transport chain complex III and complex I inhibitors, respectively, are used to inhibit mitochondrial respiration and reveal the amount non-mitochondrial respiration that contributes to baseline OCR.

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Fig. 1 Mitochondrial Stress Test. Neonatal rat ventricular myocytes were isolated and plated at a density of 30,000 cells per well in an XF96 Cell Culture Microplate in complete growth medium for 24 h. After this period, the Mitochondrial Stress Test was performed in the presence of 5.5 mM glucose and 200 μM palmitate-BSA. One micromolar oligomycin (Oligo), 3 μM FCCP, and 1 μM/1 μM antimycin A (AA)/rotenone (Rot) were used. The results are plotted as oxygen consumption rate (OCR) (pmol/min) versus time (minutes). These data represent the mean of 12 wells. Error bars represent SEM. The bioenergetics parameters that are assessed using the Mitochondrial Stress Test are shown (as described in the Data Analysis section)

Similarly, in the Mitochondrial Coupling Assay [4–8], ADP and these inhibitors allow for examination of the mitochondrial respiration states [9]. First, OCR is measured in the presence of metabolic substrate (State 2 respiration). Addition of ADP then stimulates State 3 respiration. Next, similar to the Mitochondrial Stress Test, subsequent additions of oligomycin and FCCP reveal State 4O and State 3u respiration, respectively. Finally, antimycin A is used to inhibit mitochondrial respiration. From these measurements the respiratory control ratio, a positive indicator of mitochondrial function, can also be calculated [2, 9]. In the following, we provide the complete protocols for these assays, as well as notes for optimization.

2 2.1

Materials All Assays

Seahorse XF Analyzer (see Note 1). Seahorse XF FluxPak (see Note 2). Oligomycin (see Note 3). FCCP. Antimycin A.

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Fig. 2 Mitochondrial Coupling Assay. Mitochondria were isolated from the hearts of adult (12 week-old), male mice and 4 μg per well were plated in an XF96 Cell Culture Microplate. The Mitochondrial Coupling Assay was then performed in the presence of 10 mM pyruvate and 2 mM malate. Four millimolar ADP, 2.5 μg/mL oligomycin (Oligo), 4 μM FCCP, and 4 μM antimycin A (AA) were used. The results are plotted as oxygen consumption rate (OCR) (pmol/min) (point-to-point) versus time (minutes). These data represent the mean of three wells. Error bars represent SEM. The respiration states that are assessed using the Mitochondrial Coupling Assay are shown (as described in the Data Analysis section)

Rotenone. DMSO. Metabolic Substrate(s) (see Note 4). Multichannel Pipette (20–200 μL volume). Reagent Reservoir. Sterile Water. pH Meter. Stir plate. 37
C Non-CO2 Incubator. 37
C Water Bath. Inverted Brightfield Microscope. 15 and 50 mL Conical Tubes. 1.5 mL Eppendorf Tubes. dH2O. Vortex. Aspirator. Sterile Bottles with 0.22 μm Filters and Caps.

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Cell Culture Medium. Seahorse XF Assay Medium (see Note 5). NaOH Solution.

2.3 Mitochondrial Coupling Assay (Permeabilized Cells)

XF Plasma Membrane Permeabilizer (PMP) Reagent. ADP. KOH Solution. 3 Mitochondrial Assay Solution (MAS): 210 mM Sucrose, 660 mM Mannitol, 30 mM KH2PO4, 15 mM MgCl2, 6 mM HEPES, 3 mM EGTA, 0.6% (w/v) Fatty Acid Free BSA.

2.4 Mitochondrial Coupling Assay (Isolated Tissue Mitochondria)

Surgical Tools for Tissue Dissection. PBS. Overhead Stirrer. Dounce Homogenizer/Tissue Grinder (Tight-fitting with Teflon Pestle). Refrigerated Microcentrifuge. Refrigerated Centrifuge with Swinging-Bucket Rotor (with Microplate Carrier Attachment). Bradford Reagent. BSA. ADP. KOH Solution. 3 Mitochondrial Assay Solution (MAS): 210 mM Sucrose, 660 mM Mannitol, 30 mM KH2PO4, 15 mM MgCl2, 6 mM HEPES, 3 mM EGTA, 0.6% (w/v) Fatty Acid Free BSA. Mitochondrial Isolation Buffer: 70 mM Sucrose, 210 mM Mannitol, 5 mM HEPES, 1 mM EGTA, 0.5% (w/v) Fatty Acid Free BSA.

3

Methods *Please note that the volumes indicated are for assays performed using the XF96e Analyzer. Volumes for assays performed using the XF24e or HS Mini and additional information can be found at https://www.agilent.com/en/product/cell-analysis/how-to-runan-assay.

3.1

All Assays

3.1.1 One or More Days Before Assay

1. Design the experiment at the Seahorse XF Analyzer using the Wave software. Use the Instrument User Guide to complete this step (see Notes 6–8).

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3.1.2 One Day Before Assay

1. Turn on Seahorse XF Analyzer and allow it to reach 37
C (the requires a minimum of 5 h). 2. Hydrate the XF Sensor Cartridge: (a) Separate the XF Sensor Cartridge (green) and the Cartridge Lid from the Utility Plate. (b) Add 200 μL of XF Calibrant Solution to each well of the Utility Plate. (c) Replace the XF Sensor Cartridge and the Cartridge Lid to the Utility Plate and ensure than the tips of the sensor probes are completely submerged in the XF Calibrant Solution. (d) Incubate the assembled XF Sensor Cartridge (with Cartridge Lid and Utility Plate) at 37
C in the non-CO2 incubator overnight (or for a minimum of 4 h). Alternative Strategy: (see Note 9) (a) Separate the XF Sensor Cartridge (green) and the Cartridge Lid from the Utility Plate. (b) Add 200 μL of sterile water to each well of the Utility Plate. (c) Replace the XF Sensor Cartridge and the Cartridge Lid to the Utility Plate and ensure than the tips of the sensor probes are completely submerged in the sterile water. (d) Incubate the assembled XF Sensor Cartridge (with Cartridge Lid and Utility Plate) at 37
C in a non-CO2 incubator overnight. (e) Aliquot 20 mL of XF Calibrant Solution into a 50 mL conical tube and incubate at 37
C in the non-CO2 incubator overnight.

3.1.3 Day of Assay

1. Hydrate the XF Sensor Cartridge (Alternative Strategy continued): (see Note 9). (a) Remove the assembled XF Sensor Cartridge and 50 mL conical tube containing XF Calibrant Solution from the non-CO2 incubator. (b) Separate the XF Sensor Cartridge (green) and the Cartridge Lid from the Utility Plate. (c) Remove sterile water from the Utility Plate and replace with 200 μL of pre-warmed XF Calibrant Solution per well. (d) Replace the XF Sensor Cartridge and the Cartridge Lid to the Utility Plate and ensure than the tips of the sensor probes are completely submerged in the XF Calibrant Solution.

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(e) Incubate the assembled XF Sensor Cartridge (with Cartridge Lid and Utility Plate) at 37
C in a non-CO2 incubator for 45–60 min. 3.2 Mitochondrial Stress Test 3.2.1 One or More Days Before Assay

1. In an XF Cell Culture Microplate, plate and culture cells in 80 μL of standard Cell Culture Medium and culture using standard conditions for that cell type (see Note 10). A minimum of four wells should receive medium alone, without cells, for background correction. 2. If desired, treat cells according to the investigator’s experimental conditions (see Note 11). 3. Prepare Inhibitor Stock Solutions: (a) Oligomycin—Prepare a 1 mM stock solution of oligomycin in DMSO. (b) FCCP—Prepare a 1 mM stock solution of FCCP in DMSO. (c) Antimycin A—Prepare a 1 mM stock solution of Antimycin A in DMSO. (d) Rotenone—Prepare a 1 mM stock solution of Rotenone in DMSO. (e) Aliquot and store Stock Solutions at 20
C. *If using the Mitochondrial Stress Test Kit (see Note 3), follow the instructions provided with the kit.

3.2.2 Day of Assay

1. Prepare Assay Medium: (a) Aliquot and supplement Seahorse XF Assay medium with metabolic substrates (see Note 4). *This will be used for dilution of the inhibitors for injection and running the assay (approximately 26 mL total). (b) Bring Seahorse XF Assay Medium with supplements to pH 7.4 and warm in a 37
C water bath. 2. Prepare Inhibitor Working Solutions: (see Note 12). (a) Oligomycin—Use 1 mM Stock Solution to prepare a 7.5 μM Working Solution. • Add 22.5 μL of 1 mM Oligomycin to 2977.5 μL Seahorse XF Assay Medium (prepared in step 1) and vortex. (b) FCCP—Use 1 mM Stock Solution to prepare an 8.5 μM Working Solution. • Add 25.5 μL of 1 mM FCCP to 2974.5 μL Seahorse XF Assay Medium (prepared in step 1) and vortex.

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(c) Antimycin A/Rotenone—Use 1 mM Stock Solutions to prepare a 9.5 μM/9.5 μM Working Solution. • Add 28.5 μL of 1 mM Antimycin A and 28.5 μL of 1 mM Rotenone to 2943 μL Seahorse XF Assay Medium (prepared in step 1) and vortex. *Note that during the run, these inhibitors will be injected into 130, 150, and 170 μL of Seahorse XF Assay Medium, respectively, thus diluting each to a final concentration of 1 μM. 3. Load XF Sensor Cartridge Injection Ports: (see Notes 13 and 14). (a) Remove the XF Sensor Cartridge (prepared above) from the non-CO2 incubator and remove lid. (b) Using a multichannel pipette and reagent reservoir: • Load 20 μL of 7.5 μM Oligomycin (prepared in step 2) to Injection Port A (top, left). • Load 20 μL of 8.5 μM FCCP (prepared in step 2) to Injection Port B (top, right). • Load 20 μL of 9.5 μM/9.5 μM μM Antimycin A/Rotenone (prepared in step 2) to Injection Port C (bottom, left). 4. Prepare Cell Plate for Assay Run: (a) Obtain the XF Cell Culture Microplate containing cells (prepared above) and the XF Seahorse Assay Medium (prepared in step 1). (b) Examine the cells under the microscope to ensure that they are healthy. (c) With a fine tip, carefully aspirate the culture medium from the wells. (d) Using a multichannel pipette and reagent reservoir, add 130 μL pre-warmed XF Seahorse Assay Medium (prepared in step 1) per well. (e) Incubate the plate at 37
C in a non-CO2 incubator for 45–60 min. 5. Run Assay: (a) In the Wave software, open the pre-designed experiment (prepared above). (b) Use the Instrument User Guide to run the assay, as details will vary depending upon the instrument and version of software being used. (c) In general:

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• Obtain the hydrated and pre-warmed XF Sensor Cartridge with inhibitors loaded into the ports (prepared in step 3). • Remove the lid and insert the cartridge into the instrument tray in the proper orientation. • The cartridge will calibrate within the machine. This will take approximately 15–30 min. • When this step is complete, the instrument will prompt the user to continue and the Utility Plate will be ejected. • Replace the Utility plate with the XF Cell Culture Microplate containing cells (prepared in step 4) and begin the run. 6. Data Analysis: (a) In the Wave software, the data are graphed in real time. This can be viewed as O2 (mmHg), pH, OCR (pmol/ min), or ECAR (pmol/min) versus time. (b) Data analysis can then be performed in the Wave software or using Seahorse Analytics, a web-based software application (see Note 15). For this, consult the User Guide that is appropriate for the software and version being used. (c) Alternatively, the raw data values can be exported from Wave or Seahorse Analytics to Excel and analyzed by the investigator. (d) The following are some standard bioenergetic measurements assessed with the Mitochondrial Stress Test and the equations used for their calculation. • Basal Respiration ¼ (Basal OCR)  (Non-Mitochondrial OCR). • ATP-Linked Respiration ¼ (Basal OCR)  (OCR following Oligomycin injection). • Proton Leak ¼ (OCR following Oligomycin injection)  (Non-Mitochondrial OCR). • Maximal Respiration (Maximal Respiratory Capacity) ¼ OCR following FCCP injection. • Reserved (Spare) Respiratory Capacity ¼ (Maximal Respiration)  (Basal Respiration). • Non-Mitochondrial Respiration ¼ OCR following Antimycin A/Rotenone injection.

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3.3 Mitochondrial Coupling Assay (Permeabilized Cells) 3.3.1 One or More Days Before Assay

1. In an XF Cell Culture Microplate, plate and culture cells in 80 μL of standard Cell Culture Medium and culture using standard conditions for that cell type (see Note 10). A minimum of four wells should receive medium alone, without cells, for background correction. 2. If desired, treat cells according to the investigator’s experimental conditions (see Note 11). 3. Prepare 3 Mitochondrial Assay Solution (MAS): (a) Dissolve the following amounts of the indicated reagents to 950 mL dH2O. (b) Bring pH to 7.4 using KOH. (c) Bring final volume to 1 L with dH2O to achieve the indicated concentrations. 210 mM sucrose

71.88 g

660 mM mannitol

120.23 g

30 mM KH2PO4

4.08 g

15 mM MgCl2

15 mL of 1 M solution

6 mM HEPES

6 mL of 1 M solution

3 mM EGTA

12 mL of 0.25 M solution

0.6% (w/v) fatty acid free BSA

6g

(d) Filter sterilize and store at 4
C. 4. Prepare Inhibitor Stock Solutions: (a) Oligomycin—Prepare a 1 mM stock solution of oligomycin in DMSO. (b) FCCP—Prepare a 1 mM stock solution of FCCP in DMSO. (c) Antimycin A—Prepare a 1 mM stock solution of Antimycin A in DMSO. (d) Rotenone—Prepare a 1 mM stock solution of Rotenone in DMSO. (e) Aliquot and store Stock Solutions at 20
C. *If using the Mitochondrial Stress Test Kit (see Note 3), follow the instructions provided with the kit. 3.3.2 Day of Assay

1. Prepare Assay Medium (1 MAS): (a) Aliquot 3 MAS and add metabolic substrates (see Note 4), ADP (4 mM final), XF PMP Reagent (1 nM final), and dH2O to obtain 1 MAS and these final concentrations (see Notes 16 and 17).

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*This will be used for dilution of the inhibitors for injection and running the assay (approximately 26 mL total). (b) Dilute 6 mL of 3 MAS into 12 mL dH2O to make 18 mL 1 MAS. This will be used as a wash buffer. (c) Warm both solutions in a 37
C water bath. 2. Prepare Inhibitor Working Solutions: (see Note 12). (a) Oligomycin—Use 1 mM Stock Solution to prepare a 7.5 μM Working Solution. • Add 22.5 μL of 1 mM Oligomycin to 2977.5 μL 1 MAS (prepared in step 1) and vortex. (b) FCCP—Use 1 mM Stock Solution to prepare an 8.5 μM Working Solution. • Add 25.5 μL of 1 mM FCCP to 2974.5 μL 1 MAS (prepared in step 1) and vortex. (c) Antimycin A/Rotenone—Use 1 mM Stock Solutions to prepare a 9.5 μM/9.5 μM Working Solution. • Add 28.5 μL of 1 mM Antimycin A and 28.5 μL of 1 mM Rotenone to 2943 μL 1 MAS (prepared in step 1) and vortex. *Note that during the run, these inhibitors will be injected into 130, 150, and 170 μL of Seahorse XF Assay Medium, respectively, thus diluting each to a final concentration of 1 μM. 3. Load XF Sensor Cartridge Injection Ports: (see Note 13). (a) Remove the XF Sensor Cartridge (prepared above) from the non-CO2 incubator and remove lid. (b) Using a multichannel pipette and reagent reservoir: • Load 20 μL of 7.5 μM Oligomycin (prepared in step 2) to Injection Port A (top, left). • Load 20 μL of 8.5 μM FCCP (prepared in step 2) to Injection Port B (top, right). • Load 20 μL of 9.5 μM/9.5 μM Antimycin A/Rotenone (prepared in step 2) to Injection Port C (bottom, left). 4. Prepare Cell Plate for Assay Run: (a) Obtain the XF Cell Culture Microplate containing cells (prepared above) and the 1 MAS (no supplements) and 1 MAS with supplements (prepared in step 1). (b) Examine the cells under the microscope to ensure that they are healthy.

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(c) With a fine tip, carefully aspirate the culture medium from the wells. (d) Using a multichannel pipette and reagent reservoir, add 150 μL pre-warmed 1 MAS (no supplements—prepared in step 1) per well. (e) With a fine tip, carefully aspirate the solution from the wells. (f) Using a multichannel pipette and reagent reservoir, add 130 μL pre-warmed 1 MAS with supplements (prepared in step 1) per well. (g) Immediately run assay. There is no incubation step at this point in this protocol. 5. Run Assay: (a) In the Wave software, open the pre-designed experiment (prepared above). Alter the Instrument Run Protocol as described in Note 7. (b) Use the Instrument User Guide to run the assay, as details will vary depending upon the instrument and version of software being used. (c) In general: • Obtain the hydrated and pre-warmed XF Sensor Cartridge with inhibitors loaded into the ports (prepared in step 3). • Remove the lid and insert the cartridge into the instrument tray in the proper orientation. • The cartridge will calibrate within the machine. This will take approximately 15–30 min. • When this step is complete, the instrument will prompt the user to continue and the Utility Plate will be ejected. • Replace the Utility plate with the XF Cell Culture Microplate containing cells (prepared in step 4) and begin the run. 6. Data Analysis: (a) In the Wave software, the data are graphed in real time. This can be viewed as O2 (mmHg), pH, OCR (pmol/ min), or ECAR (pmol/min) versus time. The resultant OCR for this assay are typically displayed as “point-topoint.” (b) Data analysis can then be performed in the Wave software or using Seahorse Analytics, a web-based software application (see Note 15). For this, consult the User Guide that is appropriate for the software and version being used.

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(c) Alternatively, the raw data values can be exported from Wave or Seahorse Analytics to Excel and analyzed by the investigator. (d) From the Mitochondrial Coupling Assay, mitochondrial respiration states can be assessed as previously described [5] and are defined below. From these the Respiratory Control Ratio (RCR), a positive indicator of mitochondrial function, can also be calculated using the equation below. More detailed discussions of mitochondrial respiration states and RCR can be found here [2, 5, 9]. • State 3 Respiration ¼ Initial OCR measurements. • State 4O Respiration ¼ OCR following Oligomycin injection. • State 3u Respiration ¼ OCR following FCCP injection. • RCR ¼ (State 3)/(State 4O) or (State 3u/State 4O). 3.4 Mitochondrial Coupling Assay (Isolated Mitochondria) 3.4.1 One or More Days Before Assay

1. Prepare Buffers: (a) 3 Mitochondrial Assay Solution (MAS): • Dissolve the following amounts of the indicated reagents to 950 mL dH2O. • Bring pH to 7.4 using KOH. • Bring final volume to 1 L with dH2O to achieve the indicated concentrations. 210 mM sucrose

71.88 g

660 mM mannitol

120.23 g

30 mM KH2PO4

4.08 g

15 mM MgCl2

15 mL of 1 M solution

6 mM HEPES

6 mL of 1 M solution

3 mM EGTA

12 mL of 0.25 M solution

0.6% (w/v) fatty acid free BSA

6g

• Filter sterilize and store at 4
C. (b) Mitochondrial Isolation Buffer: • Dissolve the following amounts of the indicated reagents to 950 mL dH2O. • Bring pH to 7.4 using KOH.

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• Bring final volume to 1 L with dH2O to achieve the indicated concentrations. 70 mM sucrose

23.96 g

210 mM mannitol

38.26 g

5 mM HEPES

5 mL of 1 M solution

1 mM EGTA

4 mL of 0.25 M solution

0.5% (w/v) fatty acid free BSA

5g

• Filter sterilize and store at 4
C. 2. Prepare Stock Solutions: (a) ADP—Prepare a 1 M stock solution of ADP in dH2O. (b) Oligomycin—Prepare a 5 mg/mL stock solution of oligomycin in DMSO. (c) FCCP—Prepare a 10 mM stock solution of FCCP in DMSO. (d) Antimycin A—Prepare a 40 mM stock solution of Antimycin A in DMSO. (e) Aliquot and store Stock Solutions at 20
C. 3.4.2 Day of Assay

1. Prepare Assay Medium (1 MAS): (a) Dilute 12 mL of 3 MAS into 24 mL dH2O to make 36 mL 1 MAS. *This will be used for dilution of the inhibitors for injection and running the assay. (b) Aliquot 26 mL of 1 MAS and supplement with metabolic substrates (see Notes 4 and 17). Aliquot 18 mL and warm in a 37
C water bath. Place remaining 8 mL on ice. (c) Aliquot Mitochondrial Isolation Buffer and chill on ice. *This will be used for mitochondrial isolation and volumes will vary based on the amount of tissue used. (d) Ensure all solutions are at pH 7.4. 2. Prepare Inhibitor Working Solutions: (see Note 12). (a) ADP—Use 1 M Stock Solution to prepare a 40 mM Working Solution. • Add 120 μL of 1 M ADP to 2880 μL 1 MAS (prepared in step 1) and vortex. (b) Oligomycin—Use 5 mg/mL Stock Solution to prepare a 25 μg/mL Working Solution. • Add 15 μL of 5 mg/mL Oligomycin to 2985 μL 1 MAS (prepared in step 1) and vortex.

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(c) FCCP—Use 10 mM Stock Solution to prepare an 40 μM Working Solution. • Add 12 μL of 10 mM FCCP to 2988 μL 1 MAS (prepared in step 1) and vortex. (d) Antimycin A—Use 40 mM Stock Solution to prepare a 40 μM Working Solution. • Add 3 μL of 40 mM Antimycin A to 2997 μL 1 MAS (prepared in step 1) and vortex. *Note that during the run, ADP, Oligomycin, FCCP, and Antimycin A will be injected into 180, 200, 222, and 246, μL of 1 MAS, thus diluting them final concentrations of 4 mM, 2.5 μg/mL, 4 μM, and 4 μM, respectively. 3. Load XF Sensor Cartridge Injection Ports: (see Note 13). (a) Remove the XF Sensor Cartridge (prepared above) from the non-CO2 incubator and remove lid. (b) Using a multichannel pipette and reagent reservoir: • Load 20 μL of 40 mM ADP (prepared in step 2) to Injection Port A (top, left). • Load 22 μL of 25 μg/mL Oligomycin (prepared in step 2) to Injection Port B (top, right). • Load 24 μL of 40 μM FCCP (prepared in step 2) to Injection Port C (bottom, left). • Load 26 μL of 40 μM Antimycin A (prepared in step 2) to Injection Port D (bottom, right). 4. Isolate and Plate Tissue Mitochondria: (see Note 18). (a) All animal procedures should be performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and animal protocols should be approved the institute’s Institutional Animal Care and Use Committee (IACUC). (b) Euthanize mice and excise tissue. (c) Wash tissue in pre-chilled PBS and place in pre-chilled Mitochondrial Isolation Buffer (approximately 1 mL buffer per 10–15 g of tissue). (d) Mince tissue, rinse with pre-chilled Mitochondrial Isolation Buffer to remove blood, and place on ice. (e) Using an overhead stirrer and tight-fitting dounce homogenizer/tissue grinder with Teflon pestle, dounce tissue at 2000 rpm for approximately 5–10 strokes (see Note 19). (f) Transfer tissue homogenates to a clean tube and centrifuge at 800  g for 10 min at 4
C.

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(g) Transfer the supernatant to a clean tube and centrifuge at 8000  g for 10 min at 4
C. (h) Remove the supernatant and gently resuspend pellet (containing mitochondria) in a minimal volume of pre-chilled Mitochondrial Isolation Buffer. Place on ice. (i) Determine total protein concentration by performing a Bradford Assay (see Note 20). (j) Using the total protein concentration, dilute mitochondria in pre-chilled 1 MAS supplemented with metabolic substrates (prepared in step 1) to achieve desired quantity of mitochondria per well (in a 25 μL volume per well) (see Note 21). (k) Using a multichannel pipette and reagent reservoir, plate 25 μL of mitochondrial suspension per well in an XF Cell Culture Microplate. A minimum of four wells should receive only 1 MAS supplemented with metabolic substrates (prepared in step 1) for background correction. Keep plate on ice. (l) Using a refrigerated centrifuge with swinging-bucket rotor with microplate carrier attachment/capabilities, centrifuge plate at 2000  g for 20 min at 4
C. (m) Once spin is complete, add 155 μL of pre-warmed 1 MAS supplemented with metabolic substrates (prepared in step 1) per well. (n) Examine mitochondria under the microscope to ensure adherence. (o) Immediately run assay. There is no incubation step at this point in this protocol. 5. Run Assay: (a) In the Wave software, open the pre-designed experiment (prepared above). Alter the Instrument Run Protocol as described in Note 8. (b) Use the Instrument User Guide to run the assay, as details will vary depending upon the instrument and version of software being used. (c) In general: • Obtain the hydrated and pre-warmed XF Sensor Cartridge with inhibitors loaded into the ports (prepared in step 3). • Remove the lid and insert the cartridge into the instrument tray in the proper orientation. • The cartridge will calibrate within the machine. This will take approximately 15–30 min.

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• When this step is complete, the instrument will prompt the user to continue and the Utility Plate will be ejected. • Replace the Utility plate with the XF Cell Culture Microplate containing mitochondria (prepared in step 4) and begin the run. 6. Data Analysis: (a) In the Wave software, the data are graphed in real time. This can be viewed as O2 (mmHg), pH, OCR (pmol/ min), or ECAR (pmol/min) versus time. The resultant OCR for this assay are typically displayed as “point-topoint.” (b) Data analysis can then be performed in the Wave software or using Seahorse Analytics, a web-based software application (see Note 15). For this, consult the User Guide that is appropriate for the software and version being used. (c) Alternatively, the raw data values can be exported from Wave or Seahorse Analytics to Excel and analyzed by the investigator. (d) From the Mitochondrial Coupling Assay, mitochondrial respiration states can be assessed as previously described [5] and are defined below. From these the Respiratory Control Ratio (RCR), a positive indicator of mitochondrial function, can also be calculated using the equation below. More detailed discussions of mitochondrial respiration states and RCR can be found here [2, 5]. • State 2 Respiration ¼ Initial OCR measurements. • State 3 Respiration ¼ OCR following ADP injection. • State 4O Respiration ¼ OCR following Oligomycin injection. • State 3u Respiration ¼ OCR following FCCP injection. • RCR ¼ (State 3)/(State 4O) or (State 3u/State 4O).

4

Notes 1. Seahorse XF Analyzers are available in HS Mini (8-well), XFe24 (24-well), or XFe96 (96-well) formats. These analyzers function identically, however, the throughput and surface area of the wells in the culture plates differ. Thus, plating densities of cells and mitochondria, as well as assay volumes must be adjusted according to the machine that is being used.

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2. Seahorse XF FluxPaks contain XF Sensor Cartridges, XF Cell Culture Microplates, and XF Calibrant Solution. These must be compatible with the machine that is being used. XF Cell Culture Microplates are polystyrene and can be purchased with poly-D-lysine coating. In general, polystyrene plates can be coated by the user, depending upon the cell types used, or to ensure adhesion of isolated mitochondria, as previously described [6]. 3. Oligomycin, FCCP, and Antimycin A/Rotenone can be purchased together as the Seahorse XF Cell Mitochondrial Stress Test Kit. 4. Various metabolic substrates can be used alone or in combination depending on the investigator’s biological question. It is important to consider the metabolic substrate(s) that are preferred by the tested cell type(s), either at baseline or under experimental conditions. When using intact cells, it is also important to consider the benefit of providing macronutrients, such as carbohydrates (i.e. glucose) or fats (i.e. palmitate), versus metabolites, such as pyruvate, in these cell-based assays. Several common metabolic substrate solutions, including glucose, pyruvate, glutamine, and palmitate-BSA solutions, can be purchased from Agilent Technologies. 5. Seahorse XF Assay Medium is produced based on either a DMEM or RPMI formulation. They do not contain bicarbonate, glucose, pyruvate, or glutamine. They can be purchased with low or no phenol red. A Seahorse XF Assay Medium Selection Guide can be found at https://www.agilent.com/ cs/library/selectionguide/public/5991-7878EN.pdf. 6. The default Instrument Run Protocol that is programmed into the Wave software is suitable for running the Mitochondrial Stress Test. It can, however, be modified by the investigator. This includes inserting or deleting commands, altering the order of commands, and altering Mix, Wait, and Measure times. 7. For experiments using permeabilized cells, it is critical to minimize the assay run time. To do so, eliminate the equilibration step and use shortened Mix, Wait, and Measure times (Mix— 0.5 min, Wait—0.5 min, Measure—2 min for the XF96e) (Mix—0.5 min, Wait—1 min, Measure—2 min for the XF24e) in the Instrument Run Protocol (programmed in the Wave software) [7]. 8. For experiments in isolated mitochondria, the following Instrument Run Protocol (programmed in the Wave software) is recommended [5], but can be optimized by the investigator. For example, to reach State 4 Respiration, the measurement

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time can be extended and/or less ADP (typically 0.125–1.0 mM) can be used. Protocol: Calibrate, Wait—10 min, Mix—1 min, Wait— 3 min, Mix—1 min, Wait—3 min, Mix—1 min, Measure— 3 min, Mix—1 min, Measure—3 min, Mix—1 min, Inject— Port A, Mix—1 min, Measure—3–6 min, Mix 1 min, Inject— Port B, Mix—1 min, Measure—3 min, Mix—1 min, Inject— Port C, Mix—1 min, Measure—3 min, Mix—1 min, Inject— Port D, Mix—1 min, Measure—3 min, End Protocol. 9. For more information regarding hydration of the XF Sensor Car tridge see https://www.agilent.com/cs/librar y/ usermanuals/public/XFe96_DAY_BEFORE_CARTRIDGE_ HYDRATION.pdf. 10. Cell density should be optimized prior to experimentation. To do so, cells should be seeded at varying densities and the Mitochondrial Stress Test should be performed. It is recommended that basal OCR is 20–160 pmol/min for the XF96e or HS Mini or 50–400 pmol/min for the XF24e. For more information see https://www.agilent.com/cs/library/ technicaloverviews/public/5991-7994EN.pdf. Additionally, Agilent Technologies provides a database containing all publications that cite Agilent products (www.agilent.com/cellreference-database). These publications can be filtered by Industry, Research Area, Cell Type, Cell Line, Analysis Platform, Assay, Species, Language, and Publication Date, and can provide a starting point for seeding density for a particular cell type. Plating volume will vary based on the Seahorse XF Analyzer used (8-, 24-, or 96-well). For more information see https://www.agilent.com/cs/library/usermanuals/public/ XFe96_DAY_BEFORE_CELL_SEEDING.pdf. 11. Optimal cellular health is critical for optimal results in cellbased assays. It is, therefore, important to visually monitor cellular health via microscopy, particularly following experimental treatment. 12. Working concentrations of Oligomycin, FCCP, and Antimycin A/Rotenone should be optimized prior to experimentation. To do so, perform the assay with multiple doses of Oligomycin, while leaving the others unchanged. Repeat for FCCP, then Antimycin A/Rotenone. Optimal concentrations will vary by cell type. This is particularly important for FCCP, which has a steep titration curve. Additionally, high concentrations of FCCP can inhibit OCR [2]. Recommended ranges for most cell types are 0.5–2.5 μM for Oligomycin, 0.125–2.0 μM for FCCP, and 0.5–1 μM for Antimycin A/Rotenone. 13. The XF Sensor Cartridge contains four Injection Ports. With the cartridge in the proper orientation (rows lettered A–H

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from top to bottom and the plate notch in the bottom left corner), the ports are laid out as follows: Port A (top, left), Port B (top, right), Port C (bottom, left), Port D (bottom, right). Using a multichannel pipette and reagent reservoir, slowly pipette solutions into their respective ports to avoid producing bubbles. Additionally, the XF Sensor Cartridges each contain two Loading Guides that can be placed on top of the cartridge in a particular orientation to expose only the ports that are being loaded. 14. The standard Mitochondrial Stress Test can also be modified such that a test compound can be injected prior to the Oligomycin, FCCP, and Antimycin A/Rotenone injections. This can be a drug, metabolic substrate, etc. If however, an investigator chooses to inject the test compound into only a portion of the wells, medium should be injected into the remaining wells. No ports that are included in the run protocol should be left empty. 15. There are several factors that can affect OCR and the bioenergetic parameters assessed using the Mitochondrial Stress Test or Mitochondrial Coupling Assays. For cell-based assays (the Mitochondrial Stress Test and the Mitochondrial Coupling Assay in permeabilized cells), cell number and mitochondrial number are critical determinants of OCR. Thus, cellular growth and proliferation rates, viability, and plating efficiency must be considered. Further, treatments that affect mitochondrial number, by inducing mitochondrial biogenesis or enhancing or reducing mitophagy for example, will also alter OCR. In these studies and those using isolated mitochondria, other factors, such as electron transport chain stoichiometry, may also impact bioenergetics. Since these factors can be altered by experimental treatment, it is possible to normalize the data obtained by the XF analyzer [4]. Following the assay, cell number can be assessed within each well via manual or automated cell counting or genomic DNA quantification. Alternatively, total cellular protein can be measured, however this can be problematic in cases of altered cellular viability or mitochondrial number, or when plates are coated using extracellular matrix proteins. These measurements can then be used for normalization within the Wave software (see User Guide). Alternatively, these measurements and others (such as quantification of mitochondrial number via measurement of mitochondrial DNA-to-genomic DNA ratio [10], or electron transport chain complex integrity via native gel electrophoresis [11]) can be assessed in parallel experiments and considered when interpreting results. For more information see https:// www.agilent.com/cs/library/technicaloverviews/public/

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Methods_and_Strategies_for_Normalizing_Tech_Over view_022118.pdf. 16. 1 nM of XF PMP Reagent is recommended for most cell types, however, optimization may be necessary. A 0.25–4 nM range is recommended for optimization. Additionally, 4 mM ADP is recommended, but may also require optimization. For ADP a 2–4 mM range is recommended. This higher level of ADP is required to maintain ADP-stimulated respiration during the measurement period [5, 7]. 17. It is important to remember to combine metabolic substrates such as pyruvate, glutamate, and palmitoyl carnitine with malate in order to effectively stimulate respiration [8]. Pyruvate should be made fresh the day of the assay. 18. The Mitochondrial Isolation protocol was adapted from methods previously described [5, 12] and optimized for mouse heart mitochondria [13]. It may require optimization for other tissues or for cells. 19. Extent of homogenization will require optimization depending upon tissue type and homogenization materials/ technique used. 20. Alternative assays can be performed to determine total protein concentration, however, the amount of time that the isolated mitochondria are kept on ice should be minimized. 21. The quantities of isolated mitochondria that are measured using the XF96e instrument are in the microgram range (0.5–5 μg) [5]. The quantity will require optimization based on the tissue type and homogenization materials/technique used. Recommended State 2 respiration rates are between 40 and 80 pmol/min. Too high of a concentration of mitochondria will result in high State 2 respiration rates and poor responses to ADP and the inhibitors [5].

Acknowledgments I thank Dr. Maha Abdellatif (Rutgers University—Newark, NJ) for the advice, discussions and support. References 1. Barbour JA, Turner N (2014) Mitochondrial stress signaling promotes cellular adaptations. Int J Cell Biol 2014:156020. https://doi.org/ 10.1155/2014/156020 2. Brand MD, Nicholls DG (2011) Assessing mitochondrial dysfunction in cells. Biochem J

435(2):297–312. https://doi.org/10.1042/ BJ20110162 3. Li Z, Graham BH (2012) Measurement of mitochondrial oxygen consumption using a Clark electrode. Methods Mol Biol 837:63– 72. https://doi.org/10.1007/978-1-61779504-6_5

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4. Divakaruni AS, Paradyse A, Ferrick DA, Murphy AN, Jastroch M (2014) Analysis and interpretation of microplate-based oxygen consumption and pH data. Methods Enzymol 547:309–354. https://doi.org/10.1016/ B978-0-12-801415-8.00016-3 5. Rogers GW, Brand MD, Petrosyan S, Ashok D, Elorza AA, Ferrick DA, Murphy AN (2011) High throughput microplate respiratory measurements using minimal quantities of isolated mitochondria. PLoS One 6(7):e21746. https://doi.org/10.1371/journal.pone. 0021746 6. Gerencser AA, Neilson A, Choi SW, Edman U, Yadava N, Oh RJ, Ferrick DA, Nicholls DG, Brand MD (2009) Quantitative microplatebased respirometry with correction for oxygen diffusion. Anal Chem 81(16):6868–6878. https://doi.org/10.1021/ac900881z 7. Divakaruni AS, Rogers GW, Murphy AN (2014) Measuring mitochondrial function in permeabilized cells using the seahorse XF analyzer or a Clark-type oxygen electrode. Curr Protoc Toxicol 60:25.2.1–25.216. https:// doi.org/10.1002/0471140856.tx2502s60 8. Salabei JK, Gibb AA, Hill BG (2014) Comprehensive measurement of respiratory activity in permeabilized cells using extracellular flux analysis. Nat Protoc 9(2):421–438. https://doi. org/10.1038/nprot.2014.018

9. Chance B, Williams GR (1956) The respiratory chain and oxidative phosphorylation. Adv Enzymol Relat Subj Biochem 17:65–134. h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / 9780470122624.ch2 10. Pfleger J, He M, Abdellatif M (2015) Mitochondrial complex II is a source of the reserve respiratory capacity that is regulated by metabolic sensors and promotes cell survival. Cell Death Dis 6:e1835. https://doi.org/10. 1038/cddis.2015.202 11. Jeon YH, He M, Austin J, Shin H, Pfleger J, Abdellatif M (2021) Adiponectin enhances the bioenergetics of cardiac myocytes via an AMPK- and succinate dehydrogenasedependent mechanism. Cell Signal 78: 109866. https://doi.org/10.1016/j.cellsig. 2020.109866 12. Schnaitman C, Greenawalt JW (1968) Enzymatic properties of the inner and outer membranes of rat liver mitochondria. J Cell Biol 38(1):158–175. https://doi.org/10.1083/ jcb.38.1.158 13. Pfleger J, Gross P, Johnson J, Carter RL, Gao E, Tilley DG, Houser SR, Koch WJ (2018) G protein-coupled receptor kinase 2 contributes to impaired fatty acid metabolism in the failing heart. J Mol Cell Cardiol 123: 108–117. https://doi.org/10.1016/j.yjmcc. 2018.08.025

Chapter 13 Monitoring Mitochondrial Morphology and Respiration in Doxorubicin-Induced Cardiomyopathy Chowdhury S. Abdullah , Richa Aishwarya , Mahboob Morshed, Naznin Sultana Remex, Sumitra Miriyala, Manikandan Panchatcharam, and Md. Shenuarin Bhuiyan Abstract Doxorubicin (DOX)-induced cardiomyopathy constitutes dose-dependent cardiac toxicity, culminating in fatal heart failure progression. Cardiac toxicity limits effective and subsequent use of DOX in chemotherapy regimens in pediatric, adult, and recurrent cancer patients. DOX-induced profound alterations in mitochondrial morphology, dynamics, and defects in mitochondrial energy metabolism in the heart comprise key stressors in DOX-induced cardiotoxicity. Hence, the discovery of novel molecular targets and therapeutics to mitigate DOX-induced mitochondrial dysfunctions are imperative. Herein, we provided two laboratory protocols to monitor DOX-induced alterations in mitochondrial morphology and respiration in isolated primary neonatal rat cardiomyocytes. Neonatal rat cardiomyocytes are extensively used to monitor signaling mechanisms regulating cardiomyopathy in vitro. Therefore, these protocols will help researchers study the effects of novel pharmacological and genetic manipulations against DOX-induced alterations in mitochondrial morphology and energy metabolism in cardiomyocytes. Key words Doxorubicin-induced cardiomyopathy, Mitochondrial morphology, Mitochondrial respiration, Oxygen consumption rates

1

Introduction Doxorubicin (Dox) is an efficacious anthracycline-based chemotherapeutic agent widely used against a variety of solid tumors, i.e., breast, gynecological, urogenital, endocrine, brain tumors, and also in leukemia [1–3]. Despite being an effective anti-cancer drug and first-line choice of chemotherapeutic agent, DOX’s use is limited due to its dose-dependent cardiotoxicity. DOX-induced cardiotoxicity causes pathological changes in the heart termed as “Doxorubicin-induced Cardiomyopathy” that underlies fatal heart failure development in adolescent and adult cancer surviving DOX recipient patients [4–6]. Nearly 26% of the patients suffer heart

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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failure who received a lifetime cumulative dose of DOX greater than 500 mg/m2 while heart failure prevalence reaches 36% in patients receiving 600 mg/m2 of cumulative dose of DOX [1, 6– 8]. Extensive studies confirmed DOX profoundly affects cardiac mitochondrial ultrastructure and respiratory functions, including mitochondrial swelling and abnormal cristae structure [9, 10], mitochondrial DNA (mtDNA) damage [11, 12], excessive reactive oxygen species generation through redox cycling at respiratory Complex I [13–15], suppression of mitochondrial oxidative phosphorylation involved respiration [16, 17], defects in fatty acid β-oxidation resulting switch in substrate metabolism [18], reduced mitochondrial membrane potential (Ψ m) culminating to the opening of mitochondrial permeability transition pore (mPTP) [9, 19]. This leads to apoptotic mediators release from mitochondria intermembrane and matrix that eventually potentiate cell demise, constituting heart failure development [9, 19, 20]. Cardiomyocytes require a continuous supply of energy in the form of ATP (adenosine triphosphate) to maintain their contraction–relaxation cycles [21, 22]. Oxidative phosphorylation at mitochondrial ETS accounts for nearly 95% of ATP supply in the heart [23, 24]. To this point, accumulating evidence underscores a homeostatic balance between mitochondrial fission and fusion processes, which is essential in maintaining mitochondrial quality control, ultrastructure, and respiratory functions [16, 25–31]. In the light of well-documented DOX-induced defects in cardiac mitochondrial fission–fusion signaling, ultrastructure, ETS functions, and respiration, there is an urgent need to discover novel molecular effectors and therapeutic agents to protect against DOX-induced mitochondrial structural and bioenergetics defect to prevent heart failure initiation and progression. Isolated primary neonatal rat cardiomyocytes (NRCs) are postmitotic, terminally differentiated cells with intrinsic beating rhythms, can be cultured for an extended period, responsive to pharmacological stimuli, and amenable for genetic manipulations for gene gain-of-function and loss-of-function studies [32– 34]. Hence, the NRCs culture technique provides an excellent preclinical research platform to study molecular underpinnings and to discover novel molecular effectors in cardiomyocytes in vitro exclusive of confounding effects, including in vivo neurohumoral regulation, extracellular matrix, autocrine, endocrine, and paracrine effects [33, 35]. Our current protocols describe two step-by-step methods to visualize mitochondrial network morphology and mitochondrial oxygen consumption rates (OCR) in Doxorubicin-treated isolated primary ventricular cardiomyocytes from neonatal rat pups [16, 28, 30, 31]. The first protocol will allow researchers to visualize the effects of pharmacological and genetic manipulations in preserving DOX-induced altered mitochondrial network morphology. The

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second protocol utilizes the Seahorse XFe24 Extracellular Flux analyzer, which offers microplate, live cell-based oxygen consumption rates (OCRs) measurement enabling offline mitochondrial bioenergetic parameters analysis and assessment. Together, these protocols will provide guidelines to the investigators to study the effects of novel genetic manipulations and pharmacological interventions to protect against DOX-induced pathological alterations in mitochondrial morphology and respirations.

2 2.1

Materials Equipment

1. Agilent Seahorse XFe24 Extracellular Flux Analyzer. 2. Nikon A1R high-speed confocal microscope. 3. Olympus CKX53 inverted cell culture microscope. 4. Thermo Scientific™ Heracell™ VIOS 160i CO2 incubator equipped with HEPA Filters. 5. Thermo Scientific™ 1300 Series Class II Biological Safety Cabinet. 6. VWR Non-CO2 Isotemp incubator. 7. Eppendorf 5810R centrifuge. 8. Two-key cell counter (202C, BAL Supply, LLC). 9. Bright-Line Hemacytometer (Z359629, Sigma-Aldrich). 10. DeNovix DS-11 FX+ Spectrophotometer/Fluorometer.

2.2

Cell Line

2.3 Reagents and Cell Culture Supplies

1. Primary Neonatal Rat Ventricular Cardiomyocytes (NRCs). 1. Doxorubicin Hydrochloride (D1515-10mg, Sigma-Aldrich). 2. DMEM High Glucose with GlutaMAX™ supplement Cell Culture Media (10566-016, Gibco). 3. Seahorse XF DMEM medium, pH 7.4 (103575-100, Agilent Technologies Inc.). 4. MEM alpha (1
) (12571048, Gibco). 5. Fetal Bovine Serum (FBS) (16000044, Gibco). 6. Antibiotic-Antimycotic (100
) (15240-062, Gibco). 7. 0.1% Gelatin in Water (07903, STEMCELL Technologies). 8. MitoTracker™ Green FM (M7514, Invitrogen). 9. Quick Start™ Bradford 1
Dye Reagent (Cat# 500-0205; Bio-Rad). 10. Nunc™ Lab-Tek™ II Two-well Chamber slides with cover (154461; Thermo Fisher).

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11. Phosphate Buffered Saline, pH 7.4 (1
PBS) (10010031, Gibco). 12. Dimethyl sulfoxide (DMSO) (D2650, Sigma-Aldrich). 13. VECTASHIELD® Hardset™ Antifade Mounting Medium (H-1400-10; Vector Laboratories). 14. Fisherfinest™ Premium Cover Glasses (24x60-1) (125485P, Fisher Scientific). 15. Seahorse XFe24 FluxPak (102340-100, Agilent Technologies, Inc.). 16. Seahorse XF Cell Mito Stress Test Kit includes Oligomycin, Carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), Rotenone/Antimycin A (103015-100, Agilent Technologies Inc.). 17. Seahorse XF Glucose solution (1 M) (103577-100, Agilent Technologies Inc.). 18. Seahorse XF Pyruvate solution (100 mM) (103578-100, Agilent Technologies Inc.).

3

Methods

3.1 Mitochondrial Network Morphology Visualization in DoxTreated Cardiomyocytes Through Confocal Fluorescence Microscopy

1. Isolate neonatal rat ventricular cardiomyocytes (NRCs) from 1 to 2 days old rat pups from timed-pregnant Sprague-Dawley rats as detailed previously [33]. 2. Count the cells (see Note 1) and seed 1
105 cells per well in gelatin-coated (see Note 2) Nunc™ Lab-Tek™ II Two-well Chamber slides with 800 μL of MEM α cell culture media supplemented with 10% v/v FBS and 1% v/v antibioticantimycotic overnight in a humidified cell culture incubator supplied with 5% CO2 and 95% air at 37  C (see Note 3). 3. Next morning, add an additional 1200 μL of MEM α media containing 10% v/v FBS and 1% v/v antibiotic-antimycotic per well of chamber slides. 4. After 24 hours of initial cell seeding, change the media to 2 mL of DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic per well of chamber slides (see Note 4). 5. After 72 hours of culture, treat NRCs for 24 h with DOX (prepare stock in DMSO at 50 mM) at 1 μM and 10 μM doses dissolved 1 mL per well of DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic. Treat vehicle group wells with the same DMSO volume as with DOX treated wells in 1 mL of

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DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic. 6. After 24 hours of treatment, load each well with 200 nM of mitochondrial membrane potential independent MitoTracker™ Green FM dye dissolved in 1 mL of DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic (see Note 5). 7. Incubate the chamber slides in an incubator supplied with 5% CO2 and 95% air at 37  C for 40 minutes. 8. Discard the media, add 1 mL of 3.7% paraformaldehyde in 1
PBS per well for 10 minutes at room temperature to fix the cardiomyocytes. 9. Discard the fixation solution gently using a micropipette and wash the cells twice with 1 mL of 1
PBS each time for 5 minutes. 10. Mount the cardiomyocytes with VECTASHIELD® Hardset™ Antifade Mounting Medium. 11. Observe and image the cardiomyocytes at high magnification to visualize MitoTracker Green stained mitochondrial networks using a confocal fluorescence microscope. We use Nikon A1R high-speed confocal microscope using a 60
oil objective (NA ¼ 1.4) to observe Nikon NIS-Elements C software to image the cardiomyocytes (see Note 6) (Fig. 1). 12. Mitochondrial networks length and width can be measured on NIH ImageJ software (v1.6.0) using “Freehand Line” drawing on captured digital micrographs [16]. 3.2 Mitochondrial Oxygen Consumption Rates Measurement and Bioenergetic Parameters Calculation in Cardiomyocytes 3.2.1 Cardiomyocytes Seeding in XF24 Cell Culture Plates and Doxorubicin Treatment

1. Isolate neonatal rat cardiomyocytes (NRCs) from 1 to 2 days old rat pups heart ventricles from timed-pregnant SpragueDawley rats as detailed. 2. Count the cardiomyocytes using a hemacytometer (see Note 1). 3. Seed 8
104 cells per well in gelatin-coated (see Note 2) XF24 cell culture plates initially in 200 μL of MEM α cell culture media containing 10% v/v FBS and 1% v/v antibioticantimycotic for 3–4 h in a humidified cell culture incubator supplied with 5% CO2 and 95% air at 37  C (see Notes 3 and 7). 4. Add only MEM α cell culture media in four wells (one well per row) in XF24 cell culture plates to use temperature and background control in Seahorse assay. 5. Add an additional 400 μL of MEM α media containing 10% v/v FBS and 1% v/v antibiotic-antimycotic per well of cardiomyocytes after initial 3–4 hours of seeding.

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Fig. 1 Monitoring mitochondrial network organization and morphology in cardiomyocytes. Representative confocal fluorescence microscope images demonstrating Dox alone (a), MitoTracker® Red alone (b), and MitoTracker® Green (c, d) labeled neonatal rat cardiomyocytes. (a) DOX (red) accumulates at subcellular locations as appeared as fluorescent dots in the cytosol and diffuse accumulation in the nucleus of cardiomyocytes following DOX-treatment (10 μM, 24 hours). (b) MitoTracker® Red staining reveals mitochondrial network (red) as appeared in a blend of fragmented, short tubular, and long tubular forms. Hence, we do not recommend the use of MitoTracker Red dyes to use in DOX-treated cells to avoid potential overlap between them in captured fluorescence images which may lead to erroneous interpretation of the data (please see Note 5 for explanation). (c, d) MitoTracker® Green staining reveals a mixture of a fragmented, short, and long tubular network of mitochondria (green) in vehicle (c) treated cardiomyocytes while DOX treatment (10 μM, 24 hours) resulted in hyperfused tubular mitochondrial network in cardiomyocytes

6. After 24 hours, change the media to 650 μL of DMEM High Glucose GlutaMAX™ medium supplemented with 2% v/v FBS and 1% v/v antibiotic-antimycotic per well of XF24 cell culture plates (see Note 8).

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7. After 72 hours of culture, treat NRCs for 24 h with Doxorubicin (DOX) (prepare stock in DMSO at 100 mg/mL) at 1 μM and 10 μM concentrations dissolved in DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic. Treat vehicle group wells with the same DMSO volume as with DOX treated wells in DMEM High Glucose GlutaMAX™ medium containing 2% v/v FBS and 1% v/v antibiotic-antimycotic. 3.2.2 Sensor Cartridge Hydration and Inhibitors Loading into the Sensor Cartridge

1. The day before the Seahorse assay, separate cartridge lid, Sensor cartridge, Hydro booster, and Utility plate provided in XFe24 FluxPak. 2. Add 1 mL of XF Calibrant to each well of the Utility plate. 3. Place the Hydro booster on top of the Utility plate. 4. Carefully place the Sensor cartridge through the Hydro Booster plate’s openings into the Utility plate, submerging the sensors in XF calibrant. 5. Cover the Sensor cartridge with the cartridge lid and place the setup in a non-CO2 humidified incubator at 37  C overnight (see Note 9). 6. The next day, remove the Cartridge lid from the top of the Sensor cartridge and Hydro booster from between the Sensor cartridge and Utility plate. 7. Prepare XF assay media by supplementing XF DMEM media with 10 mM Glucose and 2 mM Pyruvate. Check and adjust (if necessary) the pH to 7.4 and sterile filter under a biological safety cabinet. 8. Prepare Oligomycin (100 μM), FCCP (100 μM), Rotenone, and Antimycin A (50 μM) stock solutions in XF assay media according to the manufacturer’s instructions (Agilent). 9. Load Oligomycin, FCCP, Rotenone, and Antimycin A diluted in XF assay medium (at 10
concentration) in three ports of Sensor cartridge to deliver sequentially 1 μM Oligomycin, 4 μM FCCP, 0.5 μM Rotenone, and 0.5 μM Antimycin A per well according to the Table 1 (see Note 10).

3.2.3 Sensor Cartridge Calibration and Running the Assay

1. Set up assay protocol in “Wave” software to measure mitochondrial Oxygen Consumption Rates (OCR). An example of the protocol is described in Table 2. 2. Place the Sensor cartridge on top of the Utility plate into the Seahorse XFe24 Analyzer for calibration. 3. When the calibration is running, remove the XF24 cell culture plate from the incubator. Gently aspirate the cell culture medium leaving 50 μL of the medium in each well.

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Table 1 Inhibitors preparation for loading to XFe24 sensor cartridges DMEM Injection media port (μL)

The volume of stock compound (μL)

Concentration at Loading on sensor ports (10
) (μM) cartridge port (μL)

Oligomycin (100 μM stock)

A

1350

150

10

56

FCCP (100 μM stock)

B

900

600

40

62

Rotenone and Antimycin A (50 μM stock)

C

1350

150

5

69

Inhibitors

Table 2 Seahorse XF24 analyzer OCR measurements protocol Steps

Measurement loops

Sensor cartridge calibration

–

Equilibration

–

Baseline measurement

2–3 cycles: Mix 3 min, Wait 2 min, Measure 3 min

Inject port A—oligomycin

–

Measure

2–3 cycles: Mix 3 min, Wait 2 min, Measure 3 min

Inject port B—FCCP

–

Measure

2–3 cycles: Mix 3 min, Wait 2 min, Measure 3 min

Inject port C—rotenone and antimycin A

–

Measure

2–3 cycles: Mix 3 min, Wait 2 min, Measure 3 min

4. Gently add 600 μL of warmed XF assay media (no glucose, pyruvate) into each well. 5. Gently aspirate the media while leaving 50 μL of the media in each well. 6. Add 450 μL of warmed XF assay media supplemented with 10 mM Glucose and 2 mM Pyruvate into each well of XF24 cell culture plates. 7. Place the plate in a humidified non-CO2 incubator at 37  C until the Sensor cartridge calibration is complete. 8. Once the calibration is completed, remove and discard the Utility plate while leaving the instrument’s Sensor cartridge. 9. Place XF24 cell culture pate on the platform and run the already set up protocol to measure mitochondrial oxygen consumption rates.

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Fig. 2 Measurement of mitochondrial respiration in DOX-treated neonatal rat cardiomyocytes. Examples of oxygen consumption rates (OCR) tracing in vehicle and DOX-treated (10 μM, 24 h) cardiomyocytes. Oligomycin (1 μM), FCCP (4 μM), Rotenone (0.5 μM) and Antimycin A (0.5 μM) were sequentially injected at indicated time points (black arrows). OCR values are represented as pmol/min/μg of protein. Numerical values indicate data points where average values from replicate wells at particular time points are analyzed and plotted for OCR profiles in vehicle and DOX-treated cardiomyocyte wells

10. Following the assay, carefully discard the assay medium, homogenize the cardiomyocytes by adding 100 μL of Cell Lytic M buffer per well on a rocker at room temperature for 15 minutes. 11. Measure protein concentrations in cell lysates using Bradford assay (Bio-Rad) according to manufacturer’s instructions to normalize OCR values per μg of protein. 3.2.4 Mitochondrial Respiratory Parameters Analysis

1. Once the assay protocol is finished, transfer all the wells’ OCR values for each time point of measurement into a spreadsheet for further calculations and data analysis (Fig. 2). 2. Subtract four empty wells’ average OCR values from treatment wells’ OCR values at each time point to account for background well readings. 3. Herein, the described protocol utilizes sequential addition of Oligomycin, FCCP, Rotenone, and Antimycin A to assess mitochondrial respirations in real-time in Seahorse XFe24 Flux Analyzer. 4. Following the measurements of baseline OCRs (sum of all physiological oxygen consumptions), Oligomycin, an ATP synthase (Complex V) inhibitor, is injected to measure ATP-linked OCRs. 5. Next, carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), an uncoupling agent to collapse the proton gradient

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Table 3 Key mitochondrial respiratory parameters calculation Parameters

Equations

ATP Turnover

Basal OCR (1,2,3)—OCR with Oligomycin (4,5,6)

Maximal Respiration

OCR with FCCP (7,8,9)—OCR with Rotenone/Antimycin A (10,11,12)

Reserve Capacity

OCR with FCCP (7,8,9)—Basal OCR (1,2,3)

State Apparent

4-[{Basal OCR (1,2,3)—Oligomycin OCR (4,5,6)}/{FCCP OCR (7,8,9)— Oligomycin OCR (4,5,6)}]

Coupling Efficiency

OCR with Oligomycin (4,5,6)/Basal OCR (1,2,3)

and uncouples the mitochondrial respiration from oxidative phosphorylation, is injected to measure maximal mitochondrial OCRs. 6. The last injection delivers a mixture of Rotenone, a Complex I inhibitor, and Antimycin A, a complex III inhibitor. This injection halts cellular mitochondrial respiration and allows to measure of nonmitochondrial respiration driven by processes outside the mitochondria. 7. Real-time OCR values normalized to μg of protein can be utilized to determine key mitochondrial respiratory parameters related to mitochondrial oxidative phosphorylation, as outlined in Table 3 based on the OCR plots in Fig. 2.

4

Notes 1. Incorporate Trypan Blue (T8154, Sigma-Aldrich) during cell counting to count viable cells for subsequent seeding into the Chamber slides and XF24 cell culture plates. 2. Please coat Nunc™ Lab-Tek™ II Two-well Chamber slides (1 mL per well) and XF24 cell culture plates (500 μL per well) with 0.1% Gelatin (STEMCELL Technologies) in a biological safety cabinet at room temperature overnight before seeding cardiomyocytes into them. This step is essential to improve homogenous cardiomyocytes settling and seeding into the wells. 3. To improve cardiomyocytes settling and seeding into the wells of two-well chamber slides and XF24 cell culture plates, please follow sequential addition of cell culture media as described (initially in a low volume of cell culture media, then the

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addition of a remaining volume of cell culture media). The initial culture period in a low volume of cell culture media allows cardiomyocytes to settle rapidly and homogenously in the wells’ bottom. 4. During aspiration of cell culture media, in all steps, please avoid using a high vacuum suction technique to aspirate cell culture media from the wells as this leads to cell dislodging and inadvertent drying of the cardiomyocytes. Instead, use a micropipette to aspirate the media or buffer and add relevant cell culture media or buffer immediately. 5. Doxorubicin (DOX) possesses intrinsic fluorescence properties with fluorescence emission maximum (λemission, max) near 600 nm, which are readily detectable with photomultipliers and CCD cameras fluorescence microscopes [36–40]. Hence, we recommend using MitoTracker® Green dye (excitation maximum: 490 nm, emission maximum: 516 nm) over MitoTracker® dyes that emission maximum lies near 600 nm (e.g., MitoTracker® Red CMXRos, λemission, max ¼ 599 nm, Catalogue Number: M7512, Invitrogen) (Fig. 1). 6. We do not recommend storing MitoTracker™ Green labeled cardiomyocyte slides long term as we observed fading of MitoTracker™ Green dye’s fluorescence intensity over time. The stained cardiomyocyte slides should be kept in the dark slide box at 4  C and imaged within 24–48 hours of staining (Fig. 1). 7. During loading the XF24 cell culture plates with cell suspension in cell culture media, do not place the pipette tip directly to the bottom of the wells. Place the pipette tip at the side of the wall of wells at a slanted angle and slowly add the cell suspension for loading. 8. During change and replenishment of cell culture media, do not entirely aspirate the media to prevent cells’ drying. Leave about 50 μL of medium to avoid cells being dried out. Be cautious and gentle to avoid inadvertent cell dislodging and damage. 9. Seahorse Sensor Cartridge hydration is a required step for its proper functioning. We recommend Seahorse Sensor Cartridge hydration in a humidified non-CO2 incubator at 37  C overnight before using in Seahorse XF Test assays. 10. Mitochondrial electron transport complex inhibitors are required to be loaded into the ports prior to calibrating the Sensor Cartridge.

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Acknowledgment This work was supported by the National Institutes of Health grants: HL122354 and HL145753 to M.S.B; HL141998 and HL141998-01S1 to SM; AA025744, AA026708, and AA025744-02S1 to MP; LSUHSC-S CCDS Finish Line Award, COVID-19 Research Award, LARC Research Award, and FeistWeiller Cancer Center IDEA Grant to M.S.B.; LSUHSC-S Malcolm Feist Cardiovascular and AHA Postdoctoral Fellowship to C.S.A. (20POST35210789) and LSUHSC-S Malcolm Feist Pre-doctoral Fellowship to R.A. We would also like to thank the COBRE research core facility. References 1. Bonadonna G, Monfardini S, De Lena M, Fossati-Bellani F (1969) Clinical evaluation of adriamycin, a new antitumour antibiotic. Br Med J 3(5669):503–506 2. Bonadonna G, Monfardini S, De Lena M, Fossati-Bellani F, Beretta G (1970) Phase I and preliminary phase II evaluation of adriamycin (NSC 123127). Cancer Res 30(10): 2572–2582 3. Sledge GW, Neuberg D, Bernardo P, Ingle JN, Martino S, Rowinsky EK, Wood WC (2003) Phase III trial of doxorubicin, paclitaxel, and the combination of doxorubicin and paclitaxel as front-line chemotherapy for metastatic breast cancer: an intergroup trial (E1193). J Clin Oncol 21(4):588–592 4. Kremer LC, van der Pal HJ, Offringa M, van Dalen EC, Voute PA (2002) Frequency and risk factors of subclinical cardiotoxicity after anthracycline therapy in children: a systematic review. Ann Oncol 13(6):819–829 5. Steinherz L, Steinherz P (1991) Delayed cardiac toxicity from anthracycline therapy. Pediatrician 18(1):49–52 6. Kajihara H, Yokozaki H, Yamahara M, Kadomoto Y, Tahara E (1986) Anthracycline induced myocardial damage. An analysis of 16 autopsy cases. Pathol Res Pract 181(4): 434–441 7. Steinherz LJ, Steinherz PG, Tan CT, Heller G, Murphy ML (1991) Cardiac toxicity 4 to 20 years after completing anthracycline therapy. JAMA 266(12):1672–1677 8. Swain SM, Whaley FS, Ewer MS (2003) Congestive heart failure in patients treated with doxorubicin: a retrospective analysis of three trials. Cancer 97(11):2869–2879 9. Dhingra R, Margulets V, Chowdhury SR, Thliveris J, Jassal D, Fernyhough P, Dorn GW

II, Kirshenbaum LA (2014) Bnip3 mediates doxorubicin-induced cardiac myocyte necrosis and mortality through changes in mitochondrial signaling. Proc Natl Acad Sci U S A 111(51):E5537–E5544 10. Hoshino A, Mita Y, Okawa Y, Ariyoshi M, Iwai-Kanai E, Ueyama T, Ikeda K, Ogata T, Matoba S (2013) Cytosolic p53 inhibits Parkin-mediated mitophagy and promotes mitochondrial dysfunction in the mouse heart. Nat Commun 4:2308 11. Palmeira CM, Serrano J, Kuehl DW, Wallace KB (1997) Preferential oxidation of cardiac mitochondrial DNA following acute intoxication with doxorubicin. Biochim Biophys Acta 1321(2):101–106 12. Serrano J, Palmeira CM, Kuehl DW, Wallace KB (1999) Cardioselective and cumulative oxidation of mitochondrial DNA following subchronic doxorubicin administration. Biochim Biophys Acta 1411(1):201–205 13. Davies KJ, Doroshow JH (1986) Redox cycling of anthracyclines by cardiac mitochondria. I. Anthracycline radical formation by NADH dehydrogenase. J Biol Chem 261(7):3060–3067 14. Davies KJ, Doroshow JH, Hochstein P (1983) Mitochondrial NADH dehydrogenasecatalyzed oxygen radical production by adriamycin, and the relative inactivity of 5-iminodaunorubicin. FEBS Lett 153(1): 227–230 15. Doroshow JH, Davies KJ (1986) Redox cycling of anthracyclines by cardiac mitochondria. II. Formation of superoxide anion, hydrogen peroxide, and hydroxyl radical. J Biol Chem 261(7):3068–3074 16. Abdullah CS, Alam S, Aishwarya R, Miriyala S, Bhuiyan MAN, Panchatcharam M, Pattillo CB,

Mitochondrial Structure and Function in DOX Cardiomyopathy Orr AW, Sadoshima J, Hill JA, Bhuiyan MS (2019) Doxorubicin-induced cardiomyopathy associated with inhibition of autophagic degradation process and defects in mitochondrial respiration. Sci Rep 9(1):2002 17. Berthiaume JM, Wallace KB (2007) Adriamycin-induced oxidative mitochondrial cardiotoxicity. Cell Biol Toxicol 23(1):15–25 18. Berthiaume JM, Wallace KB (2007) Persistent alterations to the gene expression profile of the heart subsequent to chronic Doxorubicin treatment. Cardiovasc Toxicol 7(3):178–191 19. Amgalan D, Garner TP, Pekson R, Jia XF, Yanamandala M, Paulino V, Liang FG, Corbalan JJ, Lee J, Chen Y, Karagiannis GS, Sanchez LR, Liang H, Narayanagari SR, Mitchell K, Lopez A, Margulets V, Scarlata M, Santulli G, Asnani A, Peterson RT, Hazan RB, Condeelis JS, Oktay MH, Steidl U, Kirshenbaum LA, Gavathiotis E, Kitsis RN (2020) A smallmolecule allosteric inhibitor of BAX protects against doxorubicin-induced cardiomyopathy. Nat Can 1(3):315–328 20. Vercesi AE, Castilho RF, Kowaltowski AJ, de Oliveira HCF, de Souza-Pinto NC, Figueira TR, Busanello ENB (2018) Mitochondrial calcium transport and the redox nature of the calcium-induced membrane permeability transition. Free Radic Biol Med 129:1–24 21. Stanley WC, Recchia FA, Lopaschuk GD (2005) Myocardial substrate metabolism in the normal and failing heart. Physiol Rev 85(3):1093–1129 22. Kolwicz SC Jr, Purohit S, Tian R (2013) Cardiac metabolism and its interactions with contraction, growth, and survival of cardiomyocytes. Circ Res 113(5):603–616 23. Doenst T, Nguyen TD, Abel ED (2013) Cardiac metabolism in heart failure: implications beyond ATP production. Circ Res 113(6): 709–724 24. Rosca MG, Tandler B, Hoppel CL (2013) Mitochondria in cardiac hypertrophy and heart failure. J Mol Cell Cardiol 55:31–41 25. Song M, Mihara K, Chen Y, Scorrano L, Dorn GW II (2015) Mitochondrial fission and fusion factors reciprocally orchestrate mitophagic culling in mouse hearts and cultured fibroblasts. Cell Metab 21(2):273–286 26. Song M, Franco A, Fleischer JA, Zhang L, Dorn GW II (2017) Abrogating mitochondrial dynamics in mouse hearts accelerates mitochondrial senescence. Cell Metab 26(6): 872–883.e875 27. Wai T, Garcia-Prieto J, Baker MJ, Merkwirth C, Benit P, Rustin P, Ruperez FJ, Barbas C, Ibanez B, Langer T (2015)

219

Imbalanced OPA1 processing and mitochondrial fragmentation cause heart failure in mice. Science 350(6265):aad0116 28. Abdullah CS, Aishwarya R, Alam S, Morshed M, Remex NS, Nitu S, Kolluru GK, Traylor J, Miriyala S, Panchatcharam M, Hartman B, King J, Bhuiyan MAN, Chandran S, Woolard MD, Yu X, Goeders NE, Dominic P, Arnold CL, Stokes K, Kevil CG, Orr AW, Bhuiyan MS (2020) Methamphetamine induces cardiomyopathy by Sigmar1 inhibition-dependent impairment of mitochondrial dynamics and function. Commun Biol 3(1):682 29. Abdullah CS, Alam S, Aishwarya R, Miriyala S, Panchatcharam M, Bhuiyan MAN, Peretik JM, Orr AW, James J, Osinska H, Robbins J, Lorenz JN, Bhuiyan MS (2018) Cardiac dysfunction in the sigma 1 receptor knockout mouse associated with impaired mitochondrial dynamics and bioenergetics. J Am Heart Assoc 7(20):e009775 30. Alam S, Abdullah CS, Aishwarya R, Miriyala S, Panchatcharam M, Peretik JM, Orr AW, James J, Robbins J, Bhuiyan MS (2018) Aberrant mitochondrial fission is maladaptive in desmin mutation-induced cardiac proteotoxicity. J Am Heart Assoc 7(14):e009289 31. Alam S, Abdullah CS, Aishwarya R, Morshed M, Nitu SS, Miriyala S, Panchatcharam M, Kevil CG, Orr AW, Bhuiyan MS (2020) Dysfunctional mitochondrial dynamic and oxidative phosphorylation precedes cardiac dysfunction in R120G-alphaBcrystallin-induced desmin-related cardiomyopathy. J Am Heart Assoc 9(23):e017195 32. Sutherland FJ, Hearse DJ (2000) The isolated blood and perfusion fluid perfused heart. Pharmacol Res 41(6):613–627 33. Razzaque MA, Robbins J (2014) Isolation of neonatal and adult rat cardiomyocytes. In: Manual of research techniques in cardiovascular medicine. John Wiley & Sons, Ltd, New York, NY, pp 117–124 34. Parameswaran S, Kumar S, Verma RS, Sharma RK (2013) Cardiomyocyte culture - an update on the in vitro cardiovascular model and future challenges. Can J Physiol Pharmacol 91(12): 985–998 35. Tachibana S, Chen C, Zhang OR, Schurr SV, Hill C, Li R, Manso AM, Zhang J, Andreyev A, Murphy AN, Ross RS, Cho Y (2019) Analyzing oxygen consumption rate in primary cultured mouse neonatal cardiomyocytes using an extracellular flux analyzer. J Vis Exp (144):59052 36. Abdullah CS, Ray P, Alam S, Kale N, Aishwarya R, Morshed M, Dutta D,

220

Chowdhury S. Abdullah et al.

Hudziak C, Banerjee SK, Mallik S, Banerjee S, Bhuiyan MS, Quadir M (2020) Chemical architecture of block copolymers differentially abrogate cardiotoxicity and maintain the anticancer efficacy of doxorubicin. Mol Pharm 17(12):4676–4690 37. Duray PH, Cuono CB, Madri JA (1986) Demonstration of cutaneous doxorubicin extravasation by rhodamine-filtered fluorescence microscopy. J Surg Oncol 31(1):21–25 38. Karukstis KK, Thompson EH, Whiles JA, Rosenfeld RJ (1998) Deciphering the

fluorescence signature of daunomycin and doxorubicin. Biophys Chem 73(3):249–263 39. Lankelma J, Dekker H, Luque FR, Luykx S, Hoekman K, van der Valk P, van Diest PJ, Pinedo HM (1999) Doxorubicin gradients in human breast cancer. Clin Cancer Res 5(7): 1703–1707 40. Shah S, Chandra A, Kaur A, Sabnis N, Lacko A, Gryczynski Z, Fudala R, Gryczynski I (2017) Fluorescence properties of doxorubicin in PBS buffer and PVA films. J Photochem Photobiol B 170:65–69

Chapter 14 Creation of Yeast Models for Evaluating the Pathogenicity of Mutations in the Human Mitochondrial Gene MT-ATP6 and Discovering Therapeutic Molecules De´borah Tribouillard-Tanvier, Alain Dautant, Franc¸ois Godard, Camille Charles, Chiranjit Panja, Jean-Paul di Rago, and Roza Kucharczyk Abstract Numerous diseases in humans have been associated with mutations of the mitochondrial genome (mtDNA). This genome encodes 13 protein subunits of complexes involved in oxidative phosphorylation (OXPHOS), a process that provides aerobic eukaryotes with the energy-rich adenosine triphosphate molecule (ATP). Mutations of the mtDNA may therefore have dramatic consequences especially in tissues and organs with high energy demand. Evaluating the pathogenicity of these mutations may be difficult because they often affect only a fraction of the numerous copies of the mitochondrial genome (up to several thousands in a single cell), which is referred to as heteroplasmy. Furthermore, due to its exposure to reactive oxygen species (ROS) produced in mitochondria, the mtDNA is prone to mutations, and some may be simply neutral polymorphisms with no detrimental consequences on human health. Another difficulty is the absence of methods for genetically transforming human mitochondria. Face to these complexities, the yeast Saccharomyces cerevisiae provides a convenient model for investigating the consequences of human mtDNA mutations in a defined genetic background. Owing to its good fermentation capacity, it can survive the loss of OXPHOS, its mitochondrial genome can be manipulated, and genetic heterogeneity in its mitochondria is unstable. Taking advantage of these unique attributes, we herein describe a method we have developed for creating yeast models of mitochondrial ATP6 gene mutations detected in patients, to determine how they impact OXPHOS. Additionally, we describe how these models can be used to discover molecules with therapeutic potential. Key words Mitochondrial diseases, MT-ATP6 gene, Yeast, Mitochondrial DNA mutations, Drug screening, ATP synthase, Mitochondrial transformation

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-2309-1_28 Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, Corrected Publication 2022

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Introduction Mitochondria support aerobic respiration and produce the bulk of cellular ATP through the process of oxidative phosphorylation (OXPHOS), which involves the oxidation of carbohydrates and fatty acids [1]. Typically, OXPHOS involves five hetero-oligomeric protein complexes (CI-V) largely anchored in the mitochondrial inner membrane. CI-IV transfer electrons to oxygen coupled to the formation of an energy-rich transmembrane proton gradient that is used by CV (ATP synthase) to produce ATP from ADP and inorganic phosphate. The OXPHOS system contains approximately 90 different protein subunits. Their genes are located in mitochondrial and nuclear DNA. As such, two separate translation machineries, one cytosolic and the other inside the mitochondrion, are utilized to synthesize their protein products [2]. Any defect in the functioning or biogenesis of the OXPHOS system will affect high-energy demanding tissues and organs and especially the brain which consumes two thirds of the dozens of kilograms of ATP that our body uses each day. Despite its small size, the mitochondrial genome has been quite frequently implicated in mitochondrial disorders (20% of cases), presumably because of its exposure to damaging reactive oxygen species (ROS) [3]. Evaluating the pathogenicity and functional consequences of mtDNA mutations in patient’s cells and tissues may be difficult. Indeed, they usually affect only a fraction of the numerous copies of the mitochondrial genome, up to 10,000 in a single cell, which is referred to as heteroplasmy (Fig. 1b). Furthermore, the high mutability of the mitochondrial genome results in numerous family or population-specific polymorphisms [4, 5], and mutations of this genome can be influenced with nucleotide changes in nuclear and mitochondrial DNA that are per se not detrimental to human health [6, 7]. Another level of complexity is the absence of methods for genetically transforming human mitochondria. The yeast Saccharomyces cerevisiae provides a convenient model to investigate human mtDNA mutations in a well-defined genetic background [8]. Owing to its good capacity to produce ATP by the fermentation of sugars, this unicellular fungus can survive mutations that inactivate oxidative phosphorylation (Fig. 1a). It is amenable to manipulation of the mitochondrial genome [9] and is unable to stably maintain heteroplasmy [10], which makes it possible to isolate strains where all the mtDNA molecules carry the same mutation (homoplasmy) (Fig. 1b). We herein describe a method for creating yeast models of mutations of the mitochondrial MT-ATP6 gene detected in patients with various disorders [11]. This gene codes for an evolutionary conserved ATP synthase subunit, called a or 6, that is essential for moving protons across the mitochondrial inner

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Fig. 1 Attributes of the yeast S. cerevisiae for the modeling of human mitochondrial DNA mutations. (a) In addition to its capacity to produce ATP by the process of oxidative phosphorylation (OXPHOS), S. cerevisiae can make ATP efficiently by the fermentation of sugars (glucose). Thus, when provided with glucose, OXPHOSdeficient mutants can be kept alive and grown, whereas they fail to proliferate from non-fermentable carbon sources like glycerol. This phenotype may be used to identify drugs improving mitochondrial function in yeast models of mitochondrial diseases (see Fig. 7). (b) Mutations of the mitochondrial genome in patient’s cells often coexist with wild-type copies of this genome, which is referred to as heteroplasmy. Mitochondria of S. cerevisiae cannot stably maintain such a genetic heterogeneity, a property that enables to isolate strains in which all the mitochondrial DNA molecules carry the same mutation (homoplasmy), which is helpful for investigating the consequences of mtDNA mutations detected in patients

membrane coupled to ATP synthesis [12, 13]. Our method uses a yeast strain (called MR10) in which the coding sequence of ATP6 is replaced with ARG8m (atp6::ARG8m). This is a mitochondrial version of a nuclear gene (ARG8) encoding a mitochondrial protein (Arg8) involved in arginine biosynthesis [14]. An equivalent of the disease mutation (atp6mut) is first introduced into the yeast ATP6 gene carried by a plasmid (pATP6). The mutated plasmid (pATP6mut) is then delivered with a biolistic system into the mitochondria of a yeast strain totally lacking mtDNA (ρ0). The resulting mitochondrial transformant (synthetic ρ
atp6mut) is crossed with MR10 (ρ+ atp6::ARG8m), to integrate through homologous DNA recombination the atp6mut mutation in a complete mitochondrial genome (ρ+ atp6mut). When ATP synthase function is not abolished, the ρ+ atp6mut strain can be positively selected owing to its ability to grow from respiratory carbon sources that require functional mitochondria to be metabolized (like glycerol). When the ATP synthase is totally inactivated by the atp6mut mutation, the mutant is identified by virtue of its incapacity to grow in the absence of an external source of arginine. For those atp6mut mutations with highly detrimental consequences, we describe a procedure for the isolation of drugs that can attenuate their effects on mitochondrial function to help development of therapeutic molecules.

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2

Materials

2.1

Yeast Strains

The genotypes and sources of yeast strains are listed in Table 1.

Table 1 Genotypes and sources of yeast strains Name

Nuclear genotype


Strains used for constructing ρ atp6

mut

and ρ atp6 +

mut

mtDNA

Source

ρ0

[14]

+

strains

DFS160 MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1 NB403C

MATa lys2 leu2-3,112 ura3-52 his3ΔHinDIII arg8::HIS3

ρ cox2-62

[14]

W3031B

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 can1-100

ρ+

[15]

MR6

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ HIS3

[16]

MR1

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

MR10

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6::ARG8m [16] HIS3

SDC30

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1




ρ atp6

mut

ρ
atp6::ARG8m [16] COX2

ρ
ATP6 COX2

[17]

strains

RKY14

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-L237R COX2

[18]

SDC31

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-L173R COX2

[17]

RKY36

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-W126R COX2

[19]

RKY37

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-L237P COX2

[20]

RKY12

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-L173P COX2

[21]

AKY13

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-S240P COX2

[22]

AKY14

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-S242P COX2

[22]

RKY104 MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-S165N COX2

[23]

RKY109 MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ
atp6-F145S COX2

[24] (continued)

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Table 1 (continued) Name

Nuclear genotype

mtDNA

Source

ρ+ atp6mut strains MR14

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-L173R HIS3

[24]

RKY39

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-W126R HIS3

[19]

RKY38

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-L237P HIS3

[20]

RKY20

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-L173P HIS3

[21]

RKY25

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-L237R HIS3

[18]

AKY5

MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-S240P HIS3

[22]

RKY66

MATα leu2Δ ura3-52 ade2-101 arg8::URA3 kar1-1

ρ+ atp6-S242P

[22]

RKY105 MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ atp6-S165N HIS3

[23]

RKY108 MATa ade2-1 his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8:: ρ+ atp6-F145S HIS3

[24]

+

2.2

Media

2.2.1 For Growing Yeast Strains

1. YPGA (rich glucose): 2 g/L Bacto yeast extract, 2 g/L Bacto Peptone, 20 g/L (2%) glucose, and 40 mg/L adenine. 2. YPGA10 (rich glucose): 2 g/L Bacto yeast extract, 2 g/L Bacto Peptone, 100 g/L (10%) glucose, and 40 mg/L adenine. 3. YPGlyA (rich glycerol): 2 g/L Bacto yeast extract, 2 g/L Bacto Peptone, 20 mL/L (2%) glycerol. 4. CSM-Leu (complete synthetic medium lacking leucine): 1.7 g/L YNB w/o (Yeast Nitrogen base without amino acids), 5 g/L ammonium sulfate, 0.8 g/L CSM-Leu drop out mix, 20 g/L glucose (2%), and 40 mg/L adenine. 5. CSM-Arg (complete synthetic medium lacking arginine): 1.7 g/L YNB w/o (Yeast Nitrogen base without amino acids), 5 g/L ammonium sulfate, 0.8 g/L CSM-Arg drop out mix, 20 g/L glucose (2%), and 40 mg/L adenine. 6. BIOL-Leu (complete synthetic medium lacking leucine for biolistic transformation): 182.5 g/L sorbitol, 1.7 g/L YNB w/o (Yeast Nitrogen base without amino acids), 5 g/L ammonium sulfate, 0.8 g/L CSM-Leu drop out mix, 50 g/L glucose (5%), and 40 mg/L adenine. BIOL-Leu medium is solidified with 5% Bacto agar, all the other media are solidified with 2% Bacto agar. The pH of

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synthetic media is adjusted to 5.6 with NaOH to avoid hydrolysis of agar during autoclaving. 2.2.2 For Growing Bacteria

2.3 Site-Directed Mutagenesis of Plasmid DNA

LBA: 10 g/L Tryptone, 10 g/L NaCl, 5 g/L Bacto yeast extract, 100 mg/L ampicillin. The pH is adjusted to 7.0 with NaOH. This medium is solidified with 2% Bacto agar. 1. Q5® Site-Directed Mutagenesis Kit (NEBioLabs) or QuikChange Multi Site-Directed Mutagenesis Kit (Agilent). 2. Highly competent bacteria (at least 107 transformants/μg of DNA). 3. Oligonucleotides for PCR mutagenesis, DNA amplification, and sequencing are listed in Table 2.

2.4 Biolistic Transformation of Yeast Cells

1. PDS-1000/He particle delivery system and items (Tungsten M-10 microcarrier, macrocarriers, macrocarrier holders and 1350 Psi rupture disks, all from Bio-Rad).

2.4.1 Equipment 2.4.2 Reagents

1. Sterile 50% glycerol (stored at room temperature). 2. 70% and 100% ethanol solutions (stored at
20  C). 3. Filter sterilized CaCl2 2.5 M (stored at 4  C). 4. Spermidine 0.2 g in 1 mL of sterile water (stored at
20  C).

2.5 Screening of Chemical Libraries

1. 144 cm2 (12 cm  12 cm) square petri dishes. 2. Sterile paper filters (0.6 cm of diameter). 3. Chemical libraries of FDA-approved compounds (see Note 1).

3

Methods Biolistic transformation of yeast mitochondria is inefficient when performed directly into yeast cells containing wild-type (ρ+) mitochondria [9]. To avoid this problem, a suitable strain totally lacking mtDNA (ρ0) is first bombarded with the plasmid borne mutagenic mitochondrial DNA fragment, a step that takes advantage of the remarkable capacity of S. cerevisiae mitochondria to replicate any type of DNA [9]. The resulting mitochondrial transformant (synthetic ρ
) is then crossed with the host strain, to integrate (by homologous DNA recombination) the mutated DNA in a complete mitochondrial genome. To ease the introduction of mutations in the ATP6 gene, we created a yeast strain (called MR10) in which the coding sequence of this gene is replaced with ARG8m. 1. Construction of plasmid pMR1 containing an ARG8m cassette flanked by the 50 - and 30 -UTR sequences of the ATP6 gene (atp6::ARG8m).

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227

Table 2 Oligonucleotides Name

Sequence

Primers for site-directed mutagenesis of ATP6 atp6L173R-F

50 -CGCTAGAGCTATTTCAAGAGGTTTAAGATTAGGTTCTAATAT CTTAGCTGG

atp6L173R-R

50 -CCAGCTAAGATATTAGAACCTAATCTTAAACCTCTTGAAATA GCTCTAGCG

atp6L173P-F

50 -GCTAGAGCTATTTCACCAGGTTTAAGATTAGG

atp6L173P-R

50 -CCTAATCTTAAACCTGGTGAAATAGTCTAGC

atp6L273R-F

50 -GGATATGTCTGGGCTATTAGAACAGCATCATATTTAAAAGAT GC

atp6L273R-R

50 -GCATCTTTTAAATATGATGCTGTTCTAATAGCCCAGACATATCCC

atp6L237P-F

50 -GGATATGTCTGGGCTATTCCAACAGCATCATATTTA

atp6L237P-R

50 -TAAATATGATGCTGTTGGAATAGCCCAGACATATCC

atp6W126R-F

50 -CTCTTTAAGTATTGTTATTAGATTAGGTAATACTATTTTAGG

atp6W126R-R

50 -CCTAAAATAGTATTACCTAATCTAATAACAATACTTAAAGAG

atp6S240P-F

50 -GTCTGGGCTATTTTAACAGCACCATATTTAAAAGATGCAGTAT ACTTACAT

atp6S240P-R

50 -ATGTAAGTATACTGCATCTTTTAAATATGGTGCTGTTAAAATA GCCCAGAC

atp6L242P-F

50 -GTCTGGGCTATTTTAACAGCATCATATCCAAAAGATGCAGTAT ACTTACAT

atp6L242P-R

50 -ATGTAAGTATACTGCATCTTTTGGATATGATGCTGTTAAAATA GCCCAGAC

atp6S175N-F

50 -CCTTTATTAGTTATTATTGAAACTTTAAATTATTTCGCTAGAGC TATTTCATTAGG

atp6S175N-R

50 -CCTAATGAAATAGCTCTAGCGAAATAATTTAAAGTTTCAATAA TAACTAATAAAGG

atp6F145S-F

50 -GGTTTATATAAACATGGTTGAGTATTCTTCTCATTATCAGTAC CTGCTGGTACACCATTACC

atp6F145S-R

50 -GGTAATGGTGTACCAGCAGGTACTGATAATGAGAAGAATACT CAACCATGTTTATATAAACC

Primers for amplification and sequencing of ATP6 ATP6-1

50 -TAATATACGGGGGTGGGTCCCTCAC

ATP6-10

50 -GGGCCGAACTCCGAAGGAGTAAG

ATP6-F

50 -GGATCCGCGGACCCCAAAGGAGGAG

ATP6-R

50 -GGATCCGGGCCGAACTCCGAAGGAGTAAG (continued)

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Table 2 (continued) Name

Sequence

Primers for amplification and cloning of atp6::ARG8m cassette ATP6-ARG8m-F

50 -GCTCTAGATAATAAGATATAATTATGATTAATTATTTATAAGTT ATAGTTTTATAAATTTATAATTATTATGACACATTTAGAAAGAAG

ATP6-ARG8m-R

50 -GCGGGATCCTTTATTATAGTTTAATACTCCATATGTAAATTATT TTATTTTATAATTTTATTTTATAATTTAAGCATATACAGCTTCG

Restriction sites in primers are underlined. In ATP6-ARG8m-F and ATP6-ARG8m-R, the sequences in bold on the 30 side enable priming on the beginning (F) and end (R) regions of ARG8m, while those on their 50 side result in amplification of sequences upstream (F) and downstream (R) of the ATP6 coding sequence (Fig. 2). In the primers for ATP6 mutagenesis the changed codon is in bold

To create a yeast strain in which the ATP6 gene is replaced by a recoded ARG8 gene (ARG8m) [16], we first PCR amplified ARG8m (carried by plasmid pDS24 [14]) with primers ATP6-ARG8m-F and ATP6-ARG8m-R (Table 2), which yields a DNA fragment where ARG8m is flanked by the 60 base pairs upstream and downstream of the ATP6 coding sequence (Fig. 2). The ATP6-ARG8m-F and ATP6-ARG8m-R primers contain XbaI and BamHI restriction sites to ease cloning of the atp6::ARG8m DNA cassette into plasmid pJM2 [14], yielding plasmid pMR1. This plasmid contains the yeast mitochondrial COX2 gene as a marker for mitochondrial transformation (see below). 2. Construction of strain MR10 in which the coding sequence of ATP6 is replaced with a mitochondrial version (ARG8m) of the nuclear gene ARG8. As a first step, the ρ0 strain DFS160 (109 cells plated on a complete synthetic medium lacking leucine (BIOL-Leu)) was bombarded with tungsten particles coated with plasmids pMR1 and YEP351 [25], using a previously described method [9] (see Fig. 3 and Note 2). YEP351 carries the yeast nuclear LEU2 gene (this plasmid is here referred to as pLEU2). DFS160 cells are leucine auxotrophic due to a LEU2 mutation (see Table 1). Once transformed with pLEU2, they will be able to grow in the absence of an external source of leucine. Usually, 500 Leu+ transformants per plate are obtained. A few of them (1%), called MR1, will additionally contain in their mitochondria the plasmid pMR1 (ρ
atp6::ARG8m). The Leu+ transformants are replicated with a sterile velvet onto a layer of 107 cells of wild-type strain MR6 (see Note 3) plated on YPGA. In the resulting zygotes, ρ
atp6::ARG8m and ρ+ WT mitochondrial DNAs are brought into contact after the fusion of MR1 and MR6 mitochondria. The 50 and 30 UTR sequences of ATP6 that

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Fig. 2 Creation of a plasmid containing an ARG8m cassette flanked by ATP6 50 - and 30 -UTRs. Using plasmid pDS24 [14] as a template, the primers ATP6-ARG8m-F and ATP6-ARG8m-R (see Table 2) were used to amplify a DNA fragment containing ARG8m flanked by 60 bps of ATP6 50 -UTR and 30 -UTR, respectively. Both the primers also contained the restriction sites permitted to clone PCR product into plasmid pJM2 [14], yielding plasmid pMR1. This plasmid also contains the ampicillin resistance AmpR gene for amplification in E. coli and the yeast mitochondrial COX2 gene that can be used as a marker of mitochondrial genetic transformation by bombardment of cells with this plasmid

flank ARG8m in ρ
atp6::ARG8m can recombine with the wildtype ATP6 locus. Due to the presence of the nuclear kar1-1 mutation in MR1, nuclear karyogamy is strongly delayed [26], which enables the emergence of haploid recombinant ρ+ atp6:: ARG8m cells with the nuclear background of strain MR6 (Fig. 3). These cells, called MR10, are selected by virtue of their capacity to grow in glucose media lacking arginine and their incapacity to grow on respiratory media (glycerol) (see Note 4).

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Fig. 3 Replacement of the ATP6 coding sequence with ARG8m in the mitochondrial DNA of the recipient strain MR6. The plasmids pMR1 and pLEU2 were delivered by bombardment into cells of strain DFS160, which is auxotrophic for leucine (leu2Δ) and arginine (arg8Δ) and devoid of mitochondrial DNA (ρ0). Among the resulting nuclear Leu+ transformants (selected on BIOL-Leu medium), a few additionally contain pMR1 in mitochondria (ρ
atp6::ARG8m). In crosses with the recipient strain MR6, the atp6::ARG8m allele can replace ATP6 in MR6 mtDNA by homologous DNA recombination. The nuclear kar1-1 mutation present in DFS160 strongly delays nuclear karyogamy [16]. After a dozen of mitotic divisions, cells (called MR10) with the MR6 nucleus and recombinant ρ+ atp6::ARG8m mtDNA with no remaining copy of WT mtDNA emerge. These cells are respiratory growth deficient (Gly-) and prototrophic for arginine (Arg+)

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3. Construction of the template vector, pATP6, for sitedirected mutagenesis of the ATP6 gene. To mutagenize the mitochondrial ATP6 gene, the wildtype ATP6 locus was PCR amplified with primers ATP6-F and ATP6-R, using the mtDNA of strain MR6 as a template as described in Fig. 4 (see Note 5). These primers both contain a BamHI restriction site (Table 2) to facilitate the cloning of the PCR product into plasmid pJM2, yielding plasmid pATP6 (Fig. 4). 4. Introduction of atp6mut mutations in plasmid pATP6. A specific mutation of ATP6 (designated by atp6mut) is introduced into plasmid pATP6 using a Site-Directed Mutagenesis Kit and two mutagenic primers (atp6mut-F and atp6mutR (Table 2)) as described in Fig. 4. After amplification and ligation of the mutated plasmid, the non-mutated pATP6 template is eliminated by DpnI digestion and Escherichia coli is transformed with the reaction mix (see Note 6). 5. Mitochondrial transformation of the ρ0 strain DFS160 with plasmid pATP6mut. A layer of 109 cells of strain DFS160 plated on BIOL-Leu medium is bombarded with plasmids pATP6mut and pLEU2. Those Leu+ transformants that contain pATP6mut in mitochondria are termed ρ
atp6mut (Fig. 5). 6. Replacement of atp6::ARG8m in strain MR10 with the mutated ATP6 gene present in strain ρ
atp6mut. The mutated ATP6 gene in ρ
atp6mut can be integrated in a complete (ρ+) mitochondrial genome by crossing with strain MR10 in which the coding sequence of ATP6 is replaced with ARG8m (ρ+ atp6::ARG8m). In the zygotes, the mitochondria of the two strains fuse and the mutated ATP6 gene can replace atp6::ARG8m by DNA homologous recombination (Fig. 5). After a dozen of mitotic divisions, ρ+ cells homoplasmic for the atp6mut allele (ρ+ atp6mut) without any remaining copy of the arginine marker emerge. Two different procedures (see below) are used to isolate the ρ+ atp6mut cells based on the influence (mild or severe) of the atp6mut mutation on ATP synthase. (a) Case of a mild atp6mut mutation. If the atp6mut mutation induces only a partial deficit in mitochondrial ATP production (C [19, 36], m.8909T>C [24], m.8969G>A [23, 37], m.8993T>G [17, 38], m.8993T>C [21], m.9176T>G [18, 39], m.9176T>C [20], m.9185T>C [22], and m.9191T>C [22, 40]. These mutations affect evolutionary conserved residues of the protein encoded by the ATP6 gene (Fig. 8a). In high resolution structure of the yeast ATP synthase [11, 12], which is almost identical to the enzyme in mammals, the mutations map in regions important for moving protons across the mitochondrial membrane (Fig. 8b). All had a significant impact on yeast ATP synthase with deficits in mitochondrial ATP production ranging from 20% to more than 90% (Fig. 8c), in correlation with the severity of the diseases induced by these mutations. A number of drugs like chlorhexidine [41], sodium pyrithione [42], and uncoupling agents [22, 40] were identified for their ability to improve mitochondrial function in both yeast and human cells carrying these mutations. We also modeled in yeast equivalents of five human MT-ATP6 mutations found in tumors [43].

4

Notes 1. FDA-approved chemical libraries can be purchased from different companies, including Prestwick Chemical, Tebu-Bio, or Tocris. Importantly, the screening assay must be performed with cells of the atp6mut mutant freshly grown in glucose to limit accumulation of genetic reversions that improve mitochondrial function, first in YPGA10 from the
80  C mutant stock (for 1 day without shaking) and then in YPGA for 4–6 h (with shaking) before the homogeneous plating of the cells (0.125 OD in 250 μL) with sterile glass beads on a square (12 cm  12 cm) YPGlyA plate. It is recommended not to use more than 30 drugs on a single plate. All the drugs are at 10 mM in DMSO (stored at
20  C in microtitration plates), and 2 μL of each is potted on the paper disks. One disk (in a corner of the plate) is spotted with 2 μL of DMSO as a negative control. The plates are incubated at 28  C (or another temperature at which the atp6mut mutation impairs respiratory growth of yeast) and scanned usually after 5–7 days.

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Fig. 8 Examples of yeast models of mutations of the mitochondrial ATP6 gene detected in patients. (a) Evolutionary conservation of residues of the protein encoded by ATP6 (subunit a) targeted by mutations found in patients (m.8851T>C, m.8909T>C, m.8969G>A, m.8993T>C, m.8993T>G, m.9176T>C, m.9176T>G, m.9185T>C, and m.9191T>C) and the amino acid changes they induce. The shown sequences are from Homo sapiens (H.s.), Bos taurus (B.t.), Arabidopsis thaliana (A.t.), Schizosaccharomyces pombe (S.p.), Podospora anserina (P.a.), Yarrowia lipolytica (Y.l.), and Saccharomyces cerevisiae (S.c.). (b) Topology of the equivalent mutations in the yeast subunit a, viewed from the mitochondrial matrix. This protein together with a ring of 10 identical c subunits move protons from the intermembrane space (IMS) to the mitochondrial matrix coupled to rotation of the c-ring and ATP synthesis. (c) Influence of the mutations on the rate of ATP synthesis in yeast mitochondria

2. The biolistic delivery of DNA into yeast cells has been described ([9] and references therein). A critical step is the binding of the DNA to metallic particles (in our hands tungsten gives much better results than gold). 5 μg (1 μL) of YEP351 (pLEU2) and 20 μg (4 μL) of the plasmid to be introduced into mitochondria (both stored in water at
20  C) are added to 100 μL of tungsten beads in 50% glycerol. After a subsequent addition of 100 μL of 2.5 M CaCl2 and 2.9 μL of spermidine 1.38 M, the suspension is incubated on

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ice for 15 min with a very gentle mixing every 3 min. The beads coated with the DNA are harvested (for 30 min at 12,000  g at 4  C), and washed two times with 200 μL of 100% ethanol kept at
20  C (the pellet, now compact, is gently crushed with a tip and 5–6 pipetting after each ethanol addition). Finally, the pellet is resuspended in 60 μL of 70% ethanol and rapidly distributed on six macrocarriers (these are kept for at least 20 min under the hood to completely evaporate the ethanol). For the shooting, we highly recommend the use of 1300 Psi rupture disks, and there is no need to use the grids proposed by Bio-Rad (these in our hands decrease significantly the number of transformants). The vacuum in the chamber where the petri dish is placed should ideally be 29 mm of mercury and be reached in a maximum of 20 s, which is easily obtained if the system is sufficiently sealed. After firing, it is important to quickly bring the system back to atmospheric pressure in order to best preserve the viability of the yeast cells. Because of the high osmolarity of the BIOL-Leu medium (1 M sorbitol), the Leu+ transformants grow slowly and take 4–5 days to make eye-visible colonies. 3. MR6 is a derivative of W303-1B strain (Table 1) obtained after (1) replacement of the nuclear ARG8 gene with HIS3 (arg8:: HIS3, abbreviated arg8Δ), (2) substitution of its mitochondrial genome by the one of S288C strain that has been entirely sequenced, and (3) correction of the can1-1 mutation to restore a wild-type CAN1 gene [16]. The CAN1 gene encodes a basic amino acid permease that is required for a good growth of arginine auxotrophic strains in synthetic or minimal media [44]. 4. It is necessary to grow (in YPGA10) cells from the MR1  MR6 cross for at least 10–15 divisions, to obtain genetically pure ρ+ atp6::ARG8m recombinant (MR10) cells without any remaining copy of the wild-type mitochondrial DNA. Phenotypically, these cells can grow in glucose medium lacking arginine (CSM-Arg), but are unable to grow from glycerol (YPGlyA). They recover respiratory competency after crossing with SDC30, which is a synthetic ρ
derivative of DFS160 strain harboring in mitochondria the plasmid pATP6 (ρ
ATP6wt [16]). It is to be noted that the mitochondrial genome is relatively unstable in MR10 strain due to the total inactivation of ATP synthase [16]. It is therefore recommended, when possible, to grow it in complete synthetic media lacking arginine (CSM-Arg) to avoid the over accumulation of ρ
/ρ0 cells in cultures of this strain. 5. Mutagenesis of plasmid pATP6 is done according to the instructions provided by the manufacturer. We use the XL1 blue or DH5α strains of E. coli for the selection and

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amplification of the mutated plasmids (on LBA plates). Usually at least 20 bacterial transformants need to be analyzed to get one with the expected mutation. This is the most difficult step in creating mitochondrial DNA mutants. 6. Because it is AT rich, PCR amplification of mitochondrial DNA is rather difficult. The Q5 polymerase from NEBioLabs (with an elongation time of 90 s/kb) and, as a template, DNA extracted from isolated mitochondria (instead of whole cells) give the best results.

Acknowledgments This work was supported by NCN grant UMO-2016/23/B/ NZ3/02098 to RK and AFM (Association Franc¸aise contre les Myopathies) grant no. 22382 to DTT. References 1. Saraste M (1999) Oxidative phosphorylation at the fin de sie`cle. Science 283(5407): 1488–1493 2. Ott M, Herrmann JM (2010) Co-translational membrane insertion of mitochondrially encoded proteins. Biochim Biophys Acta 1803(6):767–775 3. Richter C (1992) Reactive oxygen and DNA damage in mitochondria. Mut Res 275(3–6): 249–255 4. Moraes CT (2001) What regulates mitochondrial DNA copy number in animal cells? Trends Genet 17(4):199–205 5. D’Souza AD, Parikh N, Kaech SM, Shadel GS (2007) Convergence of multiple signaling pathways is required to coordinately up-regulate mtDNA and mitochondrial biogenesis during T cell activation. Mitochondrion 7(6):374–385 6. Cai W, Fu Q, Zhou X, Qu J, Tong Y, Guan MX (2008) Mitochondrial variants may influence the phenotypic manifestation of Leber’s hereditary optic neuropathy-associated ND4 G11778A mutation. J Genet Genomics 35(11):649–655 7. Swalwell H, Blakely EL, Sutton R, Tonska K, Elstner M, He L, Taivassalo T, Burns DK, Turnbull DM, Haller RG, Davidson MM, Taylor RW (2008) A homoplasmic mtDNA variant can influence the phenotype of the pathogenic m.7472Cins MTTS1 mutation: are two mutations better than one? Eur J Hum Genet 16(10):1265–1274

8. Lasserre JP, Dautant A, Aiyar RS, Kucharczyk R, Glatigny A, TribouillardTanvier D, Rytka J, Blondel M, Skoczen N, Reynier P, Pitayu L, Rotig A, Delahodde A, Steinmetz LM, Dujardin G, Procaccio V, di Rago JP (2015) Yeast as a system for modeling mitochondrial disease mechanisms and discovering therapies. Dis Model Mech 8(6): 509–526 9. Bonnefoy N, Fox TD (2001) Genetic transformation of Saccharomyces cerevisiae mitochondria. Methods Cell Biol 65:381–396 10. Okamoto K, Perlman PS, Butow RA (1998) The sorting of mitochondrial DNA and mitochondrial proteins in zygotes: preferential transmission of mitochondrial DNA to the medial bud. J Cell Biol 142(3):613–623 11. Dautant A, Meier T, Hahn A, TribouillardTanvier D, di Rago JP, Kucharczyk R (2018) ATP synthase diseases of mitochondrial genetic origin. Front Physiol 9:329 12. Guo H, Bueler SA, Rubinstein JL (2017) Atomic model for the dimeric FO region of mitochondrial ATP synthase. Science 358(6365):936–940 13. Srivastava AP, Luo M, Zhou W, Symersky J, Bai D, Chambers MG, Faraldo-Gomez JD, Liao M, Mueller DM (2018) High-resolution cryo-EM analysis of the yeast ATP synthase in a lipid membrane. Science 360(6389):eaas9699 14. Steele DF, Butler CA, Fox TD (1996) Expression of a recoded nuclear gene inserted into yeast mitochondrial DNA is limited by

Yeast Pathogenic mtDNA Mutation Models mRNA-specific translational activation. Proc Natl Acad Sci U S A 93(11):5253–5257 15. Schacherer J, Ruderfer DM, Gresham D, Dolinski K, Botstein D, Kruglyak L (2007) Genome-wide analysis of nucleotide-level variation in commonly used Saccharomyces cerevisiae strains. PLoS One 2(3):e322 16. Rak M, Tetaud E, Godard F, Sagot I, Salin B, Duvezin-Caubet S, Slonimski PP, Rytka J, di Rago JP (2007) Yeast cells lacking the mitochondrial gene encoding the ATP synthase subunit 6 exhibit a selective loss of complex IV and unusual mitochondrial morphology. J Biol Chem 282(15):10853–10864 17. Rak M, Tetaud E, Duvezin-Caubet S, Ezkurdia N, Bietenhader M, Rytka J, di Rago JP (2007) A yeast model of the neurogenic ataxia retinitis pigmentosa (NARP) T8993G mutation in the mitochondrial ATP synthase6 gene. J Biol Chem 282(47):34039–34047 18. Kucharczyk R, Salin B, di Rago JP (2009) Introducing the human Leigh syndrome mutation T9176G into Saccharomyces cerevisiae mitochondrial DNA leads to severe defects in the incorporation of Atp6p into the ATP synthase and in the mitochondrial morphology. Hum Mol Genet 18(15):2889–2898 19. Kucharczyk R, Giraud MF, Brethes D, Wysocka-Kapcinska M, Ezkurdia N, Salin B, Velours J, Camougrand N, Haraux F, di Rago JP (2013) Defining the pathogenesis of human mtDNA mutations using a yeast model: the case of T8851C. Int J Biochem Cell Biol 45(1):130–140 20. Kucharczyk R, Ezkurdia N, Couplan E, Procaccio V, Ackerman SH, Blondel M, di Rago JP (2010) Consequences of the pathogenic T9176C mutation of human mitochondrial DNA on yeast mitochondrial ATP synthase. Biochim Biophys Acta 1797(6–7): 1105–1112 21. Kucharczyk R, Rak M, di Rago JP (2009) Biochemical consequences in yeast of the human mitochondrial DNA 8993T>C mutation in the ATPase6 gene found in NARP/MILS patients. Biochim Biophys Acta 1793(5): 817–824 22. Kabala AM, Lasserre JP, Ackerman SH, di Rago JP, Kucharczyk R (2014) Defining the impact on yeast ATP synthase of two pathogenic human mitochondrial DNA mutations, T9185C and T9191C. Biochimie 100: 200–206 23. Wen S, Niedzwiecka K, Zhao W, Xu S, Liang S, Zhu X, Xie H, Tribouillard-Tanvier D, Giraud MF, Zeng C, Dautant A, Kucharczyk R, Liu Z, di Rago JP, Chen H (2016) Identification of G8969>A in mitochondrial ATP6 gene that

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severely compromises ATP synthase function in a patient with IgA nephropathy. Sci Rep 6: 36313 24. Ding Q, Kucharczyk R, Zhao W, Dautant A, Xu S, Niedzwiecka K, Su X, Giraud MF, Gombeau K, Zhang M, Xie H, Zeng C, Bouhier M, di Rago JP, Liu Z, TribouillardTanvier D, Chen H (2020) Case report: identification of a novel variant (m.8909T>C) of human mitochondrial ATP6 gene and its functional consequences on yeast ATP synthase. Life (Basel) 10(9):215 25. Hill JE, Myers AM, Koerner TJ, Tzagoloff A (1986) Yeast/E. coli shuttle vectors with multiple unique restriction sites. Yeast 2(3): 163–167 26. Conde J, Fink GR (1976) A mutant of Saccharomyces cerevisiae defective for nuclear fusion. Proc Natl Acad Sci U S A 73(10):3651–3655 27. Emaus RK, Grunwald R, Lemasters JJ (1986) Rhodamine 123 as a probe of transmembrane potential in isolated rat-liver mitochondria: spectral and metabolic properties. Biochim Biophys Acta 850(3):436–448 28. Guerin B, Labbe P, Somlo M (1979) Preparation of yeast mitochondria (Saccharomyces cerevisiae) with good P/O and respiratory control ratios. Methods Enzymol 55:149–159 29. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227(5259): 680–685 30. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193(1):265–275 31. Rigoulet M, Guerin B (1979) Phosphate transport and ATP synthesis in yeast mitochondria: effect of a new inhibitor: the tribenzylphosphate. FEBS Lett 102(1):18–22 32. Somlo M (1968) Induction and repression of mitochondrial ATPase in yeast. Eur J Biochem 5(2):276–284 33. Bonnefoy N, Fox TD (2000) In vivo analysis of mutated initiation codons in the mitochondrial COX2 gene of Saccharomyces cerevisiae fused to the reporter gene ARG8m reveals lack of downstream reinitiation. Mol Gen Genet 262(6):1036–1046 34. di Rago J-P, Rak M, Kucharczyk R, Tetaud E, Duvezin-Caubet S (2007) Modelling in yeast of the mitochondrial ATP6 gene mutations responsible for NARP syndrome in humans and uses thereof for screening for medicaments in Patent WO/2007/125225, PCT/FR2007/000757 35. Schwimmer C, Rak M, Lefebvre-Legendre L, Duvezin-Caubet S, Plane G, di Rago JP (2006)

242

De´borah Tribouillard-Tanvier et al.

Yeast models of human mitochondrial diseases: from molecular mechanisms to drug screening. Biotechnol J 1(3):270–281 36. Kucharczyk R, Dautant A, Gombeau K, Godard F, Tribouillard-Tanvier D, di Rago JP (2019) The pathogenic MT-ATP6 m.8851T>C mutation prevents proton movements within the n-side hydrophilic cleft of the membrane domain of ATP synthase. Biochim Biophys Acta Bioenerg 1860(7):562–572 37. Skoczen N, Dautant A, Binko K, Godard F, Bouhier M, Su X, Lasserre JP, Giraud MF, Tribouillard-Tanvier D, Chen H, di Rago JP, Kucharczyk R (2018) Molecular basis of diseases caused by the mtDNA mutation m.8969G>A in the subunit a of ATP synthase. Biochim Biophys Acta Bioenerg 1859(8): 602–611 38. Su X, Dautant A, Rak M, Godard F, Ezkurdia N, Bouhier M, Bietenhader M, Mueller DM, Kucharczyk R, di Rago JP, Tribouillard-Tanvier D (2021) The pathogenic m.8993 T > G mutation in mitochondrial ATP6 gene prevents proton release from the subunit c-ring rotor of ATP synthase. Hum Mol Genet 30:381 39. Kucharczyk R, Dautant A, Godard F, Tribouillard-Tanvier D, di Rago JP (2019) Functional investigation of an universally conserved leucine residue in subunit a of ATP synthase targeted by the pathogenic m.9176T>G mutation. Biochim Biophys Acta Bioenerg 1860(1):52–59

40. Su X, Dautant A, Godard F, Bouhier M, Zoladek T, Kucharczyk R, di Rago JP, Tribouillard-Tanvier D (2020) Molecular basis of the pathogenic mechanism induced by the m.9191T>C mutation in mitochondrial ATP6 gene. Int J Mol Sci 21(14):5083 41. Couplan E, Aiyar RS, Kucharczyk R, Kabala A, Ezkurdia N, Gagneur J, St Onge RP, Salin B, Soubigou F, Le Cann M, Steinmetz LM, di Rago JP, Blondel M (2011) A yeast-based assay identifies drugs active against human mitochondrial disorders. Proc Natl Acad Sci U S A 108(29):11989–11994 42. Aiyar RS, Bohnert M, Duvezin-Caubet S, Voisset C, Gagneur J, Fritsch ES, Couplan E, von der Malsburg K, Funaya C, Soubigou F, Courtin F, Suresh S, Kucharczyk R, Evrard J, Antony C, St Onge RP, Blondel M, di Rago JP, van der Laan M, Steinmetz LM (2014) Mitochondrial protein sorting as a therapeutic target for ATP synthase disorders. Nat Commun 5:5585 43. Niedzwiecka K, Kabala AM, Lasserre JP, Tribouillard-Tanvier D, Golik P, Dautant A, di Rago JP, Kucharczyk R (2016) Yeast models of mutations in the mitochondrial ATP6 gene found in human cancer cells. Mitochondrion 29:7–17 44. Grenson M, Mousset M, Wiame JM, Bechet J (1966) Multiplicity of the amino acid permeases in Saccharomyces cerevisiae. I. Evidence for a specific argininetransporting system. Biochim Biophys Acta 127(2):325–338

Chapter 15 In Vivo Analysis of Mitochondrial Protein Synthesis in Saccharomyces cerevisiae Mitochondrial tRNA Mutants Arianna Montanari Abstract I describe here a protocol for the analysis of mitochondrial protein synthesis as a useful tool to characterize the mitochondrial defects associated with mutations in mitochondrial tRNA genes. The yeast Saccharomyces cerevisiae mutants, bearing human equivalent pathogenic mutations, were used as a simple model for analysis. The mitochondrial proteins were labeled by L[35S]-methionine incorporation in growing cells, extracted from purified mitochondria, and fractionated by SDS-polyacrylamide gel electrophoresis followed by autoradiography. By this method, it is possible to distinguish different protein synthesis profiles in the analyzed mitochondrial tRNA mutants. Key words Saccharomyces cerevisiae, Mitochondria, Mitochondrial tRNA mutants, Human equivalent mutations, Mitochondrial protein synthesis, In vivo L[35S]-methionine labeling

1

Introduction Mitochondrial diseases are a group of clinically heterogeneous disorders that occur when the mitochondria functionality is compromised. Mutations in mitochondrial (mt) tRNA genes are responsible for serious health impairment. Although the tRNA genes represent about 5% of the whole human mitochondrial genome, around 70% of the pathogenic mutations have been identified in these genes (mitomap.org). Unfortunately, to date there is still no effective treatment for these pathologies [1]. Over the years, the use of models to study the mitochondrial mutations has been optimized. The usefulness of resorting to simple organisms such as yeast Saccharomyces cerevisiae is possible thanks to the extremely genetic conservation with human and correspondence with several cellular pathways. The biolistic transformation of yeast mitochondria [2] was resulted a suitable procedure to specifically introduce mutations in mt-tRNA genes equivalent to human pathogenic substitutions. The obtained yeast

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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mt-tRNA mutants have been routinely used and have become a powerful tool for analyzing mitochondrial mutations and the associated defects [3–8]. The yeast mutants can be characterized by analysis of growth and respiration capability on different carbon sources, to identify mitochondrial metabolism defects. The mitochondrial network rearrangements can also be investigated by different staining. The effect of the mutations on mt-tRNA molecules can be various, such as structural instability and alteration of secondary or ternary structure; these defects can alter the interaction capability of the mt-tRNAs with other factors of mitochondrial translation [4–9]. By Northern blot experiments, it is possible to identify the presence and the correct aminoacylability of the mutated mt-tRNAs. The mutation LeuA14G, equivalent to severe human mutation m.3243A>G [10], produces aminoacylation defect in yeast mt-tRNALeu [6]. This mutation is also associated with complete mitochondrial activity failure, severe alteration of mitochondrial morphology, and loss of mtDNA [3]. The mutant ValC25T, bearing the equivalent human mutation m.1624C>T [11], shows mild respiratory defects and structurally instability of mutated mt-tRNAVal [5, 7]. Mutations in tRNA-encoding genes can lead to defects in protein synthesis that cause severe defects of oxidative phosphorylation (OXPHOS), being some components of electron transport chain mitochondrially encoded. The S. cerevisiae mitochondrial genome contains genes coding for seven subunits of chain reaction complexes: gene CYTB for cytochrome b (Complex III); three genes OXI1, OXI2, and OXI3 for subunits CoxII, CoxIII, and CoxI, respectively, of cytochrome c oxidase (Complex IV); three genes for subunits Atp6, Atp8, and Atp9 of the ATPase (complex V), and one gene for the ribosomal protein Var1. I report here a protocol that allows to in vivo analyzing the mitochondrial translation of yeast mutants with different severity of mitochondrial defects. The effect of mutations LeuA14G and ValC25T on mitochondrial protein synthesis were here investigated. The mitochondrial proteins were labeled with L[35S]-methionine in growing cells and then extracted from purified mitochondria. The proteins were separated by SDS-polyacrylamide gel electrophoresis (PAGE) followed by autoradiography. This method makes possible to detect variable alterations in the mitochondrial protein synthesis of the analyzed mt-tRNA mutants. The results comprise missing or complete alteration in translation products, as well as the presence of variant polypeptides in the mitochondrial translation profile. In this chapter, several suggestions are also proposed by Notes to investigate the effect of mt-tRNA mutations on mitochondrial protein synthesis in yeast cells, and it may contribute to understand the molecular mechanism underlying the different pathological phenotypes.

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Materials

2.1 Growth Media and Buffers for Labeling Reaction

1. YPDlow and YPGal growth media: 1% Bacto yeast extract, 1% Bacto peptone with low glucose (0.25%) for YPDlow, or 2% galactose for YPGal. All components were dissolved in H2O and sterilized by autoclaving. 2. DAPI solution: 40 -6-Diamidino-2-phenylindole (DAPI) dissolved in H2O to final concentration of 1 μg/mL. 3. Labeling buffer: 40 mM Na-phosphate buffer pH 7.4 and 0.45% glucose. It can be sterilized by filtering and stored at room temperature (RT). 4. Cycloheximide: powder of cycloheximide dissolved in H2O to final concentration of 4 mg/mL. 5. Protoplasting buffer: 1.2 M sorbitol, 50 mM Na-phosphate pH 7.5, 1% β-mercaptoethanol, 0.2% Zymolase 20 T. This buffer can be prepared without β-mercaptoethanol and Zymolase, sterilized by filtering and stored at 20  C. The β-mercaptoethanol and Zymolase were added before being used. 6. Lysis buffer: 10 mM Tris–acetate pH 7.5, 0.25 M mannitol, 1 mM EDTA pH 7.5. This buffer can be sterilized by filtering and stored at 20  C. 7. 2 Laemmli buffer: 0.125 M Tris–HCl pH 6.8, 4% SDS, 10% β-mercaptoethanol, 20% glycerol, 0.004% bromophenol blue. It can be prepared in advance and stored at 20  C in aliquots to avoid excessive thawing of the sample.

2.2

SDS-PAGE Gel

1. 30% Acrylamide/bis-Acrylamide solution (ratio 29:1): 30 g acrylamide and 0.8 g bis-acrylamide in 100 mL H2O. 2. 12% Resolving gel solution: the reagents and specific quantities are indicated in Table 1. 3. 4% Stacking gel solution: the reagents and specific quantities are indicated in Table 2. 4. 10% APS (stock solution): ammonium persulfate (APS) dissolved in H2O at final concentration of 10%. This solution can be stored at 20  C in aliquots. Failure in gel polymerization may depend on the alteration of the stock solution. 5. 10 Running buffer (stock solution): tris(hydroxymethyl)aminomethane (Tris) 30 g, glycine 144 g, and sodium dodecyl sulfate (SDS) 10 g dissolved in H2O to have 1 L solution. The pH of the stock solution should be adjusted with HCl at pH 8.3, and it can be stored at RT. The Running buffer was used as 1 solution in the running chamber. 6. Fixing solution: 7.5% glacial acetic acid and 25% methanol.

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Table 1 List of reagents with the related amounts to prepare the Resolving gel 12% Resolving solution gel (20 mL) Volume

Stock solution

Final concentration

29:1/Acrylamide:bis-Acrylamide

8 mL

30%

12%

Tris–HCl pH 8.8

5 mL

1.5 M

375 mM

SDS

200 μL

10%

0.1%

40 μL

–

5%

200 μL

10%

0.1%

a

TEMED APS

a

H2O

Up to 20 mL

a

TEMED (N-N-N-N-tetramethylethylendiamine) and APS were added immediately before pouring the gel in order to trigger the polymerization

Table 2 List of reagents with the related amounts to prepare the Stacking gel 4% Stacking solution gel (6 mL) Volume

Stock solution

Final concentration

29:1/Acrylamide:bis-Acrylamide

800 μL

30%

4%

Tris–HCl pH 6.8

1.5 mL

0.5 M

125 mM

SDS

60 μL

10%

0.1%

TEMEDa

12 μL

–

5%

a

60 μL

10%

0.1%

APS

H2O

Up to 6 mL

a

TEMED (N-N-N-N-tetramethylethylendiamine) and APS were added immediately before pouring the gel in order to trigger the polymerization

7. Coomassie blue staining solution: 0.1% Coomassie, 10% glacial acetic acid, and 50% ethanol. 8. Destaining solution: 10% glacial acetic acid and 12.5% isopropanol.

3

Methods Mitochondrial translation products were in vivo labeled by this procedure. For treatment, cells from fresh culture were inoculated into 100-mL flask.

In Vivo Labelled Mitochondrial Proteins

3.1 Cells Preparation and In Vivo Labeling of Mitochondrial Proteins

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1. Inoculate the cell culture in 25 mL of YPDlow or YPGal media (see Note 1). 2. Grow the cells overnight at 28  C by shaking in order to collect the cells at the exponential phase (spectrophotometer absorbance OD 600 nm between 0.2 and 0.5) (see Note 2). 3. Before collecting culture for labeling, analyze the mtDNA presence by DAPI staining (see Note 3): add 1% formaldehyde to 100 μL of cell culture and after 30 min at RT, wash the cells with 100 μL of H2O. Suspend the pellet in 5 μL of DAPI solution, and after 5 min of incubation, observe the cells by fluorescence microscope at the wavelength of 350 nm. 4. Collect the cells in 50-mL tube by centrifugation at 1110  g RT for 10 min in order to have 30 mg of cells for the labeling reaction. 5. Wash the cells twice with sterile H2O and collect the cells by centrifugation at 1110  g at RT for 5 min. 6. Suspend the cells in 1 mL of Labeling buffer. For the labeling reaction 0.850 mL of cell suspension should be transferred in a 10-mL tube, containing 0.1 mL cycloheximide (4 mg/mL), and incubate at 28  C for 10 min by shaking (see Note 4). 7. Label the mitochondrial proteins by adding to the culture 4 μL (0.16 mCi) of L[35S]-methionine (40 mCi/mL) (see Note 5). The labeling reaction was stopped after 90 min adding 0.1 mL of L-methionine (0.2 M).

3.2 Purification of Mitochondria and Mitochondrial Proteins Extraction

1. Centrifuge the sample at 1110  g for 10 min at RT and wash with H2O to remove the L[35S]-methionine and cycloheximide. Transfer the cells in 1.5-mL tube by centrifugation at 900  g for 10 min at RT. 2. Suspend the cells in 1 mL of Protoplasting buffer and incubate at 37  C for 30 min. 3. Collect the protoplasts by centrifugation at 3500  g for 10 min at RT and suspend them in 1 mL of Lysis buffer. 4. Collect the mitochondria after 20 min of centrifugation at 16,200  g at +4  C. Purified mitochondria were obtained after washing twice with the Lysis buffer and centrifugation at 16,200  g at +4  C for 15 min. 5. Suspend the purified mitochondria in 50 μL of 2 Laemmli buffer. The mitochondrial proteins were extracted by vigorous pipetting of the sample and by 10 min of incubation at 95  C. The sample can be stored at 20  C after preparation and before SDS-PAGE procedure.

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3.3 SDS-Polyacrylamide Gel Electrophoresis (PAGE)

1. To separate the labeled mitochondrial proteins, the 12% SDS polyacrylamide gel should be prepared. For the gel preparation two glass plates with dimensions 20 cm  23 cm and 0.4 mm of thickness should be used. 2. Wash the glasses with 96% ethanol and with H2O twice, to remove the ethanol; dry with lint-free paper. Place the glasses in the apparatus laterally separated by the spacers (see Note 6). 3. Introduce the Resolving solution in the space between the two glass plates; the Resolving gel will be placed at the bottom. Cover with 96% ethanol to minimize the distortion of the gel surface. 4. After around 15 min of polymerization, remove the ethanol, dry the glasses with Whatman paper and pour the Stacking gel on the top. Close the gel with the comb to obtain the wells for the samples. Allow the gel polymerization for at least 1 h (see Note 7). 5. Attach the two glass plates containing the gel to the electrophoresis tank of the running apparatus. Fill the upper and lower reservoirs with 1 Running buffer, making sure that the glasses were correctly positioned and that there was no loss of buffer from the tanks. 6. Incubate the samples for 5 min at 95  C before loading. Remove the comb from the top of the gel, releasing the wells for loading (see Note 8). Load 10 μL of each samples into the wells. The running was performed at +4  C, starting from 80 mA to facilitate the entrance of the samples into the wells, followed by running at constant current of 30 mA for 3 or 4 h until the blue dye reached the bottom of the gel (see Note 9).

3.4 Gel Preparation for Autoradiography

1. After the electrophoresis, separate the two glasses and wash the gel twice with H2O; remove the Stacking gel on the top, and cut sideways the parts of Resolving gel not containing samples. 2. Fix the gel with 20 min of incubation in Fixing solution RT (see Notes 10 and 11). 3. Transfer the gel onto the Whatman paper and cover it with plastic wrap. 4. Dry the gel in gel-dryer apparatus for 1 h at 80  C. After drying, cool the gel 10 min RT before analysis by autoradiography. 5. For autoradiography expose the gel for 24–48 h to X-Ray film at 80  C (see Note 12).

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4

249

Notes 1. To obtain a higher amount of mitochondria from the extraction, the choice of growth medium is not negligible, and the metabolism of yeast S. cerevisiae should be taken into account. This yeast is a facultative aerobic organism so it can grow in fermentative carbon source (such as glucose) or in respirative medium (such as glycerol as carbon source). Moreover, the carbon metabolism of this yeast is regulated by the Crabtree effect: in high glucose, the respirative pathway is reversible repressed. In intermediate condition, such as low glucose or galactose, the glucose repression is minimized and the respirative metabolism is active. In these media, the cells show an extensive mitochondrial metabolism, and it is possible to optimize the mitochondrial mass extraction. The use of low glucose or galactose should be a rational choice in particular for the mitochondrial defective mutants for which the yield extraction of mitochondria may not be considerable. 2. Yeast mitochondrial mutants, carrying mutations in different mt-tRNA genes, show defects in mitochondrial protein synthesis with different severity. It is known that inhibition of mitochondrial protein synthesis increases the formation of petite mutants during growth [12]. These mutants do not respire and can be easily identified because they produce small-sized colonies in glucose containing solid medium (Fig. 1). Colonies of normal size contain rho+ cells (with a wild-type mtDNA); compared to these, the petite colonies contain cells carrying partial deletions of mtDNA (rho) or mtDNA-less cells (rho ). Different mt-tRNA mutants showed variable percentage of petite mutants [8, 13, 14]. This phenotype is generally related to the severity of the mitochondrial defect, and it is a useful tool to characterize the effect of the mutations on the mitochondrial functionality. It might consider collecting some large colonies from a fresh plate and refresh them in liquid medium for few hours instead of grow them by overnight incubation. This alternative procedure should be chosen to avoid proceeding with the labeling of predominantly rho/rho cells. 3. The analysis of mtDNA content of the cells is necessary before proceeding with treatment. It is important to evaluate the ratio between rho+ and rho/rho cells in the sample; if most of the cells have lost their mtDNA during the growth incubation, they are not useful for mitochondrial activity study. A very rapid analysis for the detection of mtDNA content is the DAPI staining; this dye is known to form fluorescent complexes with natural double-stranded DNA and permits to visualize the mtDNA in addition to the nuclear DNA (Fig. 2).

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Fig. 1 Growth of yeast mt-tRNA mutant on glucose plate. The petite colonies containing rho/rho cells are distinguishable by small size from those containing the rho+ cells. Some of petite colonies are indicated by the arrows

Fig. 2 Fluorescence microscopy of S. cerevisiae cells stained by DAPI. In all cells, the nuclear DNA is the bigger blue dot. The cells identified by one asterisk are rho+ and their mtDNA is shown as small dots. The cell indicated by two asterisks is rho , and it does not contain mtDNA. The bar corresponds to 1 μm

4. The cycloheximide is necessary to inhibit the cytosolic translation and to enrich the sample of mitochondrial proteins. Cycloheximide is a citoplasmic protein synthesis inhibitor in eukaryotes; it inhibits the elongation step of translation

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through binding to the E-site of the ribosome and interfering with deacetylated tRNA interaction [15]. Adding the cycloheximide permits to incorporate the L[35S]-methionine only in the new synthetized mitochondrial proteins. 5. The half-life of L[35S]-methionine is 87.4 days. Despite it should be recommended using it within 1 month of delivery. 6. The two glass plates are typically one slightly different in size from the other, and spacers are used to form a watertight seal to put the unpolymerized gel solution. The glass plates were washed carefully, and the spacers were arranged at each side parallel to the two edges. The entire length of the two sides and the bottom of the plates were bound with yellow electrical tape to avoid the leakage of the gel solution. Before the electrophoresis, the tape was removed. 7. The gel could be prepared the day before the electrophoresis. In this case, the gel in the glass plates can be stored to +4  C, covered by paper wet with H2O and transparent film. This should prevent the excessive gel dehydration. 8. When the comb was removed, the wells could be dirty by the presence of gel residues; this could cause bad sample loading. For this reason, it is necessary to clean the wells by spraying the Running buffer inside with a syringe until all residues were removed. 9. To facilitate the sample loading, it is possible to perform the first steps of electrophoresis RT: loading the samples, waiting the entrance of the samples into the wells, and then transferring the electrophoretic apparatus to cold room for the running. 10. If the gel is small in size, it is possible to perform a slight shaking incubation so that the buffer was evenly distributed. In the other case, the gel could break so it is preferable to not shake. After the fixing, the Coomassie staining may be performed before the autoradiography, as control, to check that the same amount of the proteins was loaded for all samples. In this case, the use of the thicker spacers of 1.5 mm should be suggested to avoid the gel breaking. However, it is necessary to punctualize that a too thick gel could break during the subsequent dehydration phase. 11. For Coomassie staining, incubate the gel 20 min in Coomassie blue dye followed by overnight incubation in Destaining solution, to remove the Coomassie. The day after, an image can be acquired by placing the gel over a diaphanoscope. 12. Figure 3 shows the mitochondrial translation products of yeast mt-tRNA mutants LeuA14G and ValC25T compared with those of wild-type strain. While in the wild-type sample (WT) the different encoded mitochondrial proteins are well

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Fig. 3 Autoradiography of the mitochondrial proteins by SDS-PAGE electrophoresis. The mitochondrial protein extracts were in vivo labeled using 35 L[ S]-methionine from wild-type strain (WT) and from the isogenic mt-tRNA mutants, LeuA14G and ValC25T, and applied to SDS-PAGE gel. Compared to the wild-type, the mitochondrial translation of the mutant LeuA14G is completely compromised; the mutant ValC25T shows a different translation profile

distinguished, the pattern of the mutant LeuA14G is completely degraded, and it is not possible to visualize the single mitochondrial subunits. In the case of mutant ValC25T, a new profile is visible: it is possible to observe decreasing signal in the region of Cytb/CoxIII and the appearance of lower bands not detectable in the wild-type sample. These new bands might represent mitochondrial translation products with amino acidic misincorporation. It is interesting to note that the analysis of yeast amino acid usage shows that the Leucine is the most abundant amino acid in the most of mitochondrial proteins (https://www.ncbi.nlm. nih.gov/). Therefore, it is not surprising that in the mutant LeuA14G, bearing aminoacylation defect, the mitochondrial translation is totally compromised. Instead, the amino acid Valine is used on average in the mitochondrial encoded proteins. The Cytochrome b is the mitochondrial subunit with the higher percentage of Valine, compared with the other mitochondrial encoded proteins; Valine is the second most

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abundant amino acid in this subunit. So it is possible to speculate that the ValC25T mutation might produce aminoacidic misincorporation defects; in this case, the major effects could be ascribed in Cytochrome b in which the Valine is more represented. The methodology here proposed contributes to molecular characterization of the mitochondrial defect of mt-tRNA mutants by using yeast S. cerevisiae as useful simple model to specifically analyze the mitochondrial protein synthesis in growing cell culture.

Acknowledgments I am grateful to Laura Frontali and Silvia Francisci for their helpful and critical support. References 1. Russell MO, Gorman GS, Lightowlers RN, Turnbull DM (2020) Mitochondrial diseases: hope for the future. Cell 181:168–188. https://doi.org/10.1016/j.cell.2020.02.051 2. Fox TD, Sanford JC, Mc Mullin TW (1988) Plasmids can stably transform yeast mitochondria lacking endogenous mitDNA. Proc Natl Acad Sci 85:7288–7292. https://doi.org/10. 1073/pnas.85.19.7288 3. Feuermann M, Francisci S, Rinaldi T et al (2003) The yeast counterparts of human “MELAS” mutations cause mitochondrial dysfunction that can be rescued by overexpression of the mitochondrial translation factor EF-Tu. EMBO Rep 4:53–58. https://doi.org/10. 1038/sj.embor.embor713 4. De Luca C, Besagni C, Frontali L et al (2006) Mutations in yeast mt tRNAs: specific and general suppression by nuclear encoded tRNA interactors. Gene 377:169–176. https://doi. org/10.1016/j.gene.2006.04.003 5. De Luca C, Zhou YF, Montanari A et al (2009) Can yeast be used to study mitochondrial diseases? Biolistic tRNA mutants for the analysis of mechanisms and suppressors. Mitochondrion 9:408–417. https://doi.org/10.1016/ j.mito.2009.07.004 6. Montanari A, Besagni C, De Luca C et al (2008) Yeast as a model of human mitochondrial tRNA base substitutions: investigation of the molecular basis of respiratory defects. RNA 14:275–283. https://doi.org/10.1261/rna. 740108

7. Montanari A, De Luca C, Frontali L, Francisci S (2010) Aminoacyl-tRNA synthetases are multivalent suppressors of defects due to human equivalent mutations in yeast mt tRNA genes. Biochim Biophys Acta 1803: 1050–1057. https://doi.org/10.1016/j. bbamcr.2010.05.003 8. Montanari A, De Luca C, Di Micco P et al (2011) Structural and functional role of bases 32 and 33 in the anticodon loop of yeast mitochondrial tRNAIle. RNA 17:1983–1996. https://doi.org/10.1261/rna.2878711 9. Francisci S, De Luca C, Oliva R et al (2005) Aminoacylation and conformational properties of yeast mitochondrial tRNA mutants with respiratory deficiency. RNA 11:914–927. https://doi.org/10.1261/rna.2260305 10. Goto Y, Nonaka I, Horai S (1990) A mutation in the tRNALeu(UUR) gene associated with the MELAS subgroup of mitochondrial encephalomyopathies. Nature 348:651–653. https://doi.org/10.1038/348651a0 11. McFarland R, Clark KM, Morris A et al (2002) Multiple neonatal deaths due to a homoplasmic mitochondrial DNA mutation. Nat Genet 30: 145–146. https://doi.org/10.1038/ng819 12. Myers AM, Pape LK, Tzagoloff A (1985) Mitochondrial protein synthesis is required for maintenance of intact mitochondria genomes in S. cerevisiae. EMBO J 4:2087–2092 13. Montanari A, Francisci S, Fazzi D’Orsi M, Bianchi MM (2014) Strain-specific nuclear genetic background differentially affects mitochondria-related phenotypes in

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Saccharomyces cerevisiae. Microbiology Open 3:288–298. https://doi.org/10.1002/ mbo3.167 14. Garrido-Maraver J, Cordero MD, Domı´nguez ˜ in I et al (2012) Screening of effective Mon pharmacological treatments for MELAS syndrome using yeasts, fibroblasts and cybrid models of the disease. Br J Pharmacol 167:

1311–1328. https://doi.org/10.1111/j. 1476-5381.2012.02086.x 15. Helinek TG, Devlin TM, Ch’ih JJ (1982) Initial inhibition and recovery of protein synthesis in cycloheximide-treated hepatocytes. Biochem Pharmacol 31:1219–1225. https://doi. org/10.1016/0006-2952(82)90007-7

Chapter 16 Visualizing Mitochondrial Importability of a Protein Using the Yeast Bi-Genomic Mitochondrial-Split-GFP Strain and an Ordinary Fluorescence Microscope Marine Hemmerle, Bruno Senger, Jean-Paul di Rago, Roza Kucharczyk, and Hubert D. Becker Abstract Proving with certainty that a GFP-tagged protein is imported inside mitochondria by visualizing its fluorescence emission with an epifluorescence microscope is currently impossible using regular GFP-tagging. This is particularly true for proteins dual localized in the cytosol and mitochondria, which have been estimated to represent up to one third of the established mitoproteomes. These proteins are usually composed of a surpassingly abundant pool of the cytosolic isoform compared to the mitochondrial isoform. As a consequence, when tagged with a regular GFP, the fluorescence emission of the cytosolic isoform will inevitably eclipse that of the mitochondrial one and prevent the detection of the mitochondrial echoform. To overcome this technical limit, we engineered a yeast strain expressing a new type of GFP called Bi-Genomic Mitochondrial-Split-GFP (BiG Mito-Split-GFP). In this strain, one moiety of the GFP is encoded by the mitochondrial DNA while the second moiety of the GFP can be tagged to any nuclearencoded protein (suspected to be dual localized or bona fide mitochondrial). By doing so, only mitochondrial proteins or echoforms of dual localized proteins, regardless of their organismal origin, trigger GFP reconstitution that can be visualized by regular fluorescence microscopy. The strength of the BiG MitoSplit-GFP system is that proof of the mitochondrial localization of a given protein rests on a simple and effortless microscopy observation. Key words Mitochondria, Localization, Dual localized, BiG Mito-Split-GFP, Living cells, Saccharomyces cerevisiae, Epifluorescence microscopy

1

Introduction Fusing green fluorescent protein (GFP) or any other fluorescent protein still remains the fastest approach to visualize, using epifluorescence or confocal microscopy, the localization of a given

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-2309-1_28 Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, Corrected Publication 2022

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protein in a given subcellular compartment and notably in mitochondria. However, a growing number of proteins studied in yeast and human display a dual cytosolic and mitochondrial localization [1, 2]. It is therefore very difficult, if not impossible, to distinguish the fluorescence of the mitochondrial pool (also called mitochondrial echoform) of such kind of GFP-tagged dual-localized protein from the cytosolic pool (cytosolic echoform) by epi- or confocal microscopy [3]. The fact that, in most cases, the proportion of the cytosolic pool largely exceeds that of the mitochondrial one dramatically accentuates this difficulty. Confirming the mitochondrial relocation of a small pool of a cytosolic protein usually necessitates obtaining highly pure mitochondria by subcellular fractionation and verifying the purity of the mitochondria using antibodies directed against subcompartment-specific markers. However, getting ultrapure mitochondria devoid of cytosolic contaminants is technically almost impossible to achieve since disrupting and isolating compartments tightly bound together in the cell via contact sites like the ER-mitochondria ERMES complex [4] or the vacuole-mitochondria vCLAMP [5] are currently very challenging and laborious. Owing to the constantly increasing number of mitochondrial echoforms in established mitoproteomes, there is a crucial need for a reliable unbiased, cost-effective, and rapid new tool that could validate the presence of a protein inside mitochondria and that also enables the specific visualization of only the mitochondrial echoform of a dual-localized protein. To this end, we re-engineered the two-fragment self-assembling Split-GFP originally designed by Cabantous and co-workers [6] and engineered a yeast strain harboring a so-called Bi-Genomic Mitochondrial-Split-GFP (BiG Mito-Split-GFP). In this strain, the gene encoding the β1-10 fragment of the Split-GFP (GFPβ1-10) was biolistically integrated [7, 8] into the mitochondrial genome and is thus only translated inside the mitochondrial matrix by mitoribosomes. On the other hand, the second Split-GFP fragment (GFPβ11) sequence can be fused to any nuclear-encoded protein that will be translated by the cytosolic translation machinery (Fig. 1). By doing so, the cytosolic echoform of GFPβ11-tagged dual-localized protein will not generate a GFP signal because it will never be in contact with the GFPβ1-10 complementing fragment which is synthesized and entrapped inside the mitochondria. On the opposite, upon mitochondrial import, the GFPβ11 fragment fused to the mitochondrial echoform will be able to interact with the mitochondrially produced GFPβ1-10 and trigger assembly of a fully functional GFP emitting a fluorescent signal that can be visualized by conventional fluorescence microscopy. To guarantee that any mitochondrial protein will generate a fluorescent signal regardless of its expression level, we did not use a single β11 to tag the nuclear-encoded gene of interest but concatenated three β11 strands (β11-chaplets or GFPβ11ch, [7]) and mutagenized both original Split-GFP fragments in order to increase their stability and self-assembly [7, 9]. The BiG Mito-Split-GFP system not

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Fig. 1 Principle of the BiG Mito-Split-GFP system. The gene of interest (GOI) encoding a dual-localized protein has been tagged at its 30 -end with the sequence of three interspaced β11 fragments of the Split-GFP (β11chaplet; GFPβ11ch). The mRNA of this GOIβ11ch gene will be translated by cytosolic ribosomes into the corresponding protein fused at its C-terminus with the β11ch tag. The pool corresponding to the cytosolic β11ch-tagged echoform stays in the cytoplasm and is thus not generating any fluorescent signal, while the mitochondrial β11ch-tagged echoform will translocate inside the mitochondrial matrix. Upon mitochondrial import, the β11ch appended to the mitochondrial echoform will bind to GFPβ1-10 fragment of the Split-GFP translated by mitoribosomes from transcripts transcribed from a mtDNAintegrated β1-10 gene. Upon interaction, both mitochondria-restricted SplitGFP fragments will reconstitute a functional GFP yielding fluorescent restricted to mitochondria that can be visualized by epifluorescence microcopy

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only allows specific and unbiased visualization of discrete pools of mitochondrial echoforms of yeast dual-localized proteins but also constitutes a suitable and reliable tool to test mitochondrial importability of proteins from other organisms like plants and human [7]. We describe herein (1) how to generate the plasmid expressing the GFPβ11ch-tagged gene of interest, (2) how to transform and grow the BiG Mito-Split-GFP strain, (3) how to visualize the mitochondrial GFP signal with an ordinary epi-fluorescence microscope, and (4) how to verify expression of the GFPβ11ch and GFPβ110 fragments using commercially available antibodies.

2

Materials

2.1 PCR and Isothermal Assembly

Description of the buffers and isothermal assembly enzymes used to fuse the GFPβ11ch sequence at the 30 -end of the gene of interest (GOI). The design of the oligonucleotide primers used for the PCR and isothermal assembly of the plasmid are described in Fig. 2. 1. Thermal Cycler (BIO-RAD)).

(C1000

TouchTM

Thermal

Cycler

2. 5 isoT master mixture (mix): 3 mL Tris–HCl pH 7.5 1 M, 150 μL MgCl2 2 M, 600 μL of dNTP mix (10 mM each), 300 μL DTT 1 M, 300 μL NAD+ 100 mM, 1.5 g of PEG-8000, and sterile ultrapure water up to 6 mL final volume. Store 320 μL aliquots in a 80  C freezer. 3. 2 reaction master mix: 320 μL 5 isoT master mix, 1.2 μL of T5 exonuclease (New England Biolabs, 10 U/μL), 20 μL Phusion DNA polymerase (ThermoFisher Scientific, 2 U/μL), 8 μL Taq DNA ligase (New England Biolabs, 40 U/μL), and water up to 800 μL. Freeze 11 μL aliquots and keep at 80  C. 4. Dry Heating Block for 1.5-mL microtubes. 2.2 BiG Mito-SplitGFP Transformation

1. BiG Mito-Split-GFP Strain (see Table 1). 2. Liquid Yeast extract Peptone Dextrose (YPD): Peptone 2% (w/v), Yeast extract 1% (w/v), Dextrose 2% (w/v). Autoclaved at 120  C, 1.2 bars for 20 min. 3. Bench top centrifuge. 4. Dry Heating Block for 1.5-mL microtubes. 5. Salmon sperm DNA (10 mg/mL). 6. Lithium acetate 1 M (no need to adjust pH). 7. Polyethylene glycol (PEG)-4000 50% (w/v). 8. Water bath. 9. Solid Synthetic Complete Dextrose without tryptophane (SCD-Trp): 0.675% Yeast Nitrogen Base (without amino acids), 2% Dextrose, amino acid (-Trp) dropout mix, 2% agar. Autoclaved at 120  C, 1.2 bars for 20 min.

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Fig. 2 Adding the GFPβ11ch tag (a) at the 50 -end of gene of interest using the isothermal assembly strategy. (a) Sequence (nucleotides and amino acids) of the GFPβ11ch tag. The β-strands (arrows) of the GFPβ11ch tag are in green and the spacers between the β11 fragments are in black. Nucleotides in red correspond to the downstream isoT tag (used for P2-for and G2-rev). Note that underlined nucleotides show EcoRI (red) and XhoI (black) restriction sites at both ends of the GFPβ11ch. (b) In silico assembly of the desired construct (p414pGPD-GOI- GFPβ11ch) showing the selection marker (bla (ampicillin resistance), yellow), the gene of interest (GOI, blue), the GFPβ11ch tag (green) with a small linker region (red), the GPD promoter (pGPD), the yeast auxotrophy marker TRP1, the E. coli origin of replication (ColE1), and the yeast centromeric origin of replication (A-C : ARS/CEN). (C) To obtain the final construct, PCR amplification is used for the GOI and the two plasmidic fragments. The template used to amplify the destination vector is a similar plasmid containing an irrelevant GOI (named xxx here). The position and the sequence of the primers is indicated. Please note that for primer

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10. Glass beads (2.5–4.5 mm in diameter). 11. Incubator. 2.3 BiG Mito-SplitGFP Growth

1. Liquid Synthetic Complete Dextrose without tryptophane (SCD-Trp): 0.675% Yeast Nitrogen Base (without amino acids), 2% Dextrose, amino acid (-Trp) dropout mix. Autoclaved at 120  C, 1.2 bars for 20 min. 2. Incubator Shaker. 3. Liquid Synthetic Complete Galactose without tryptophane (SCGal-Trp): 0.675% Yeast Nitrogen Base (without amino acids), 2% Galactose, amino acid (-Trp) dropout mix. Do not autoclave, but rather use vacuum filtration system with 0.2 μm pores.

2.4 Epi-Fluorescence Microscopy

1. AXIO Observer d1 (Carl Zeiss) epifluorescence microscope using a 100 plan apochromatic objective (Carl Zeiss). 2. 76  26 mm microscope slide and 22  22 mm cover slip. 3. MitoTracker™ Red CMXRos (Thermo Fisher). 4. Liquid SC-Gal: 0.675% Yeast Nitrogen Base (without amino acids), 2% Galactose, amino acid (-Trp) dropout mix. Do not autoclave, but rather use vacuum filtration system with 0.2 μm pores.

2.5 SDS-PAGE Electrophoresis, Western Blotting, and Immunodetection

1. NaOH 0.185 M. 2. Trichloroacetic acid 100% (w/v). 3. Laemmli buffer 1 (see Note 1). 4. Mini-PROTEAN® 3 System Glass Plates (BIO-RAD). 5. SDS polyacrylamide gels. The resolving gel is composed of acrylamide:bisacrylamide (30%, 37.5:1) supplemented with 0.5% (v/v) 2,2,2-Trichloroethanol (TCE) (optional, see Note 2) diluted in 450 mM Tris–HCl pH 8.8, 0.12% (w/v) SDS, and the acrylamide concentration is adjusted depending on the molecular weight of the proteins to analyze (usually 8, 10, or 12% (v/v)). After addition of 0.1% (w/v) ammonium persulfate (APS) and 0.1% (v/v) N,N,N0 ,N-Tetramethylethylenediamine (TEMED), immediately pour the solution between two 10  8 cm Mini-PROTEAN® Spacer Plates from BIO-RAD with integrated spacers of 0.75, 1, or 1.5 mm. Cover the

ä Fig. 2 (continued) G2-rev, the reverse-complement sequence of your GOI has to be considered (without stop codon) since the primer has the same sequence than the minus strand of the GOI. (d) Gibson assembly of the three PCR products (the GOI and the two plasmid halves). This assembly is performed at 50  C and has three steps: (1) T5 exonuclease (50 ! 30 ) creates short 30 overhangs since it gets rapidly inactivated at 50  C, (2) the single stranded overhangs can hybridize and become substrates for Phusion DNA polymerase, and (3) the fragments are ligated together by Taq DNA ligase to yield the final product

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Table 1 BiG Mito-Split-GFP strain genotypes Strain

Nuclear genotype

Mitochondria DNA

BiG MitoSplit-GFP

MATa his3-11,15 trp1-1 leu2-3,112 ura3-1 CAN1 arg8::HIS3

ρ+ atp6::GFPβ1-10 50 UTRCOX2 ATP6 30 UTRCOX2

Table 2 Primary antibodies used in this study Specificity Name

Type

Vendor

Dilution

GFPβ1-10

Anti-GFP N-terminal

Rabbit polyclonal

Sigma (#G1544)

1:5000

GFPβ11ch

Anti-GFP

Mouse monoclonal IgG1κ clones 7.1 and 13.1

Roche 1:5000 (#11814460001)

Table 3 Secondary antibodies used in this study Targeted primary antibody

Type

Dilution

Anti-GFP N-terminal

Goat-Anti-rabbit-HRP

1:5000

Anti-GFP

Goat-Anti-mouse-HRP

1:5000

surface with ethanol 100% and discard the ethanol after polymerization. The stacking gel, which is poured on top of the resolving gel, is composed of 5% (v/v) acrylamide:bisacrylamide (30%, 37.5:1) supplemented with TCE diluted in 125 mM Tris–HCl pH 6.8, 0.1% (w/v) SDS. The polymerization is triggered upon the addition of 0.1% APS and 0.1% TEMED, and the comb is added right after pouring the gel. 6. Mini-PROTEAN® Tetra Vertical Electrophoresis Cell from BIO-RAD. 7. 0.2 μm PVDF membranes. 8. Trans-Blot Transfer Packs (BIO-RAD). 9. Trans-Blot Turbo™ transfer system from BIO-RAD. 10. Ethanol 100%. 11. TBS-Tween20: 50 mM Tris–HCl pH 7.6, 150 mM NaCl and 0.3% (v/v) Tween20. 12. Skim milk powder. 13. Heidolph Duomax shaker. 14. Primary antibody (see Table 2). 15. Secondary antibody (see Table 3).

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16. Clarity Western ECL substrate from BIO-RAD. 17. Chemidoc™ imaging system from BIO-RAD. 18. Red Ponceau.

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Methods

3.1 Engineering the Plasmid Expressing the GFPβ11ch-Tagged Gene of Interest Using the Gibson Assembly Procedure

3.2 Transforming the BiG Mito-Split-GFP Strain

1. PCR-amplify the gene of interest using the G1-for and G2-rev primer (see Fig. 2 for details). 2. The isothermal assembly is performed as follows: 11 μL 2 reaction master mix is added to 9 μL DNA solution that is classically obtained by mixing PCR fragments (1–5 fragments, approximately 1 μL each) and sterile ultrapure water. The reaction mixture is immediately incubated at 50  C for 30–60 min. Parental plasmid DNA present within PCR products can be eliminated via DpnI digestion (see Note 3). 5 μL of the whole mixture is used to transform chemically competent Escherichia coli cells [10] and yields classically 10–800 colonies. 1. Inoculate BiG Mito-Split-GFP Strain from a YPD-rich medium plate in 3 mL liquid YPD medium and grow overnight (ON) at 30  C under 200 rpm rotational shaking. 2. Pellet cells by centrifugation at 5000  g for 5 minutes (min) at room temperature (RT). 3. Heat the salmon sperm DNA (10 mg/mL) at 95  C for 5 min to solubilize it. 4. Wash cells with 1 mL sterile ultrapure water and centrifuge at 5000  g for 5 min at RT. 5. Repeat step 4. 6. Resuspend the pellet in the appropriate volume of sterile ultrapure water (see Note 4). 7. In a 1.5-mL microtube, add 10 μL of cooled salmon sperm DNA before adding 50 μL of cell suspension (see Note 5). Mix by vortexing for 5 s. 8. Add 1 μg of the p414-pGPD-GOI-GFPβ11ch (see Fig. 2) to the cells and then 72 μL sterile ultrapure water, 36 μL lithium acetate 1 M, and 240 μL polyethylene glycol (PEG)-4000 50% (w/v). Mix by vortexing for 5 s. 9. Incubate cells in a 42  C water bath for 20–40 min. 10. Harvest cells by short spin centrifugation (approximately 10 s). Add 1 mL sterile ultrapure water (see Note 6), mix by inversion without resuspending the pellet and immediately remove the supernatant to leave 100–150 μL liquid in the tube.

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11. Resuspend the pellet using the remaining supernatant. 12. Spread cells on a petri dish containing solid SCD-Trp medium using sterile glass beads. 13. Incubate the plate at 30  C for 3–4 days until colonies appear. 14. Streak colonies on a new SCD-Trp plate and incubate at 30  C for 2–3 days. 15. Select the clones that can grow on SCD-Trp and store the SCD-Trp plate at 4  C. 3.3 Culture of the Transformed BiG MitoSplit-GFP Strain

1. Grow 3–4 selected clones overnight (ON) in 3 mL liquid SCD-Trp medium at 30  C under 200 rpm rotational shaking. 2. In the morning, dilute the cells to an OD600nm ¼ 0.4 in 3 mL SC Galactose (SCGal)-Trp medium and incubate them at 30  C under 200 rpm rotational shaking to an OD600nm ¼ 0.8  1.2. 3. Separate equally the whole culture into two 1.5-mL sterile microtubes, one for the preparation of total protein extract and Western blot (to verify the expression of the GFPβ11chtagged protein; see Subheading 3.5) and the other one for mitochondria staining and microscopy imaging. 4. Centrifuge the cells for total protein extract at 5000  g for 5 min at RT and discard the supernatant. 5. Keep the cell pellet for total protein extract on ice and proceed with mitochondria staining and microscope imaging.

3.4 Visualizing Mitochondrial Importability of the GFPβ11ch-Tagged Protein

1. Add MitoTracker™ Red CMXRos (Thermo Fisher) to a final concentration of 100 μM to the cells prepared at Subheading 3.3, step 3 for mitochondria staining. 2. Incubate the cells at 30  C for 15 min. 3. Centrifuge the cells at 5000  g for 5 min at RT. 4. Wash cells with 1 mL sterile deionized water. 5. Repeat step 4 twice. 6. Resuspend cells in 50 μL SCGal medium. 7. Drop 2.3 μL of stained cells between 76  26 mm microscope slide and 22  22 mm cover slip (see Note 7). 8. Perform epifluorescence imaging (see Fig. 3) using an AXIO Observer d1 (Carl Zeiss) epifluorescence microscope using a 100 plan apochromatic objective (Carl Zeiss). The images are acquired using the GFP, TRITC, and Nomarski filters and using the camera CoolSnap HQ2 photometrix (Roper Scientific). For image processing and montage, the software ImageJ is used.

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Fig. 3 Fluorescence emission patterns of a dual-localized protein β11ch tagged and expressed in the BiG MitoSplit-GFP to that of the same protein tagged with regular GFP. Micrographs of the RS453 strain expressing GUS1 tagged with regular GFP (top) and of the BiG Mito-Split-GFP strain expressing GUS1 tagged with β11ch (bottom). GUS1 encodes glutamyl-tRNA synthetase which has been shown to be dual-localized both in the cytosol and the mitochondria [11]. Cells were treated as described in Subheading 3.4, steps 1–7 and images were taken as described in Subheading 3.4, step 8 and processed with the ImageJ software. Mitochondrial echoforms of glutamyl-tRNA synthetase are indicated with white arrowheads 3.5 Verifying Expression of the GFPβ11ch-Tagged Protein and/or of the Mitochondrial GFPβ1-10 Fragment

1. Resuspend the pellet in 450 μL NaOH 0.185 M. 2. Incubate the suspension for 10 min on ice. 3. Add 50 μL trichloroacetic acid 100%. 4. Incubate the suspension for 10 min on ice. 5. Pellet the precipitated proteins at 13,000  g for 15 min at 4  C. 6. Remove the supernatant (see Note 8) and resuspend the pellet in 75 μL Laemmli buffer 1 (see Note 9). 7. Prepare polyacrylamide gels and load 7.5 μL of total protein extract in Laemmli buffer onto the polyacrylamide gel and perform the electrophoresis at 180 V for 10 min and then at 200 V using Mini-PROTEAN® Tetra Vertical Electrophoresis Cell from BIO-RAD. 8. If TCE was added to the polyacrylamide gel, proteins having migrated can be detected using Stain-free procedure on Chemidoc™ imaging system from BIO-RAD (see Note 2).

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9. Transfer proteins onto 0.2 μm PVDF membranes using TransBlot Transfer Packs and Trans-Blot Turbo™ transfer system from BIO-RAD (see Note 10). 10. Shortly incubate the membrane in ethanol 100% and wash with 10 mL TBS-Tween20 solution. 11. Block the membrane with 10 mL TBS-Tween20 solution supplemented with 5% (w/v) skim milk for 30 min at RT under shaking on Heidolph Duomax 1030 at 10 rpm. Discard the blocking solution. 12. Incubate the membrane overnight at 4  C under shaking with 10 mL blocking solution containing the primary antibody (see Table 2) diluted 1:5000 (see Note 11). 13. The next day, remove the primary antibody solution. 14. Wash the membrane with 10 mL TBS-Tween20 solution for 10 min at RT under shaking. Discard the washing solution. 15. Repeat step 14 two times. 16. Add 10 mL of the secondary antibody (see Table 2) diluted 1: 5000 in blocking solution and incubate for 3 h at RT under shaking. 17. Remove the secondary antibody solution. 18. Wash the membrane with 10 mL TBS-Tween20 solution for 10 min at RT under shaking. 19. Repeat step 18 twice. 20. Wash the membrane with 10 mL TBS 1 solution. 21. Add the Clarity Western ECL substrate from BIO-RAD (750 μL of each reagent for one membrane). 22. Perform immunodetection using the Chemidoc™ imaging system from BIO-RAD and the Chemiluminescence program (manual acquisition and optimal time exposure). 23. Rinse the membrane with TBS-Tween20 solution shortly and add undiluted Red Ponceau staining to the membrane (see Note 12). Incubate 20 min at RT under shaking. 24. Remove the Red Ponceau staining solution and wash the membrane with 10 mL TBS-Tween20 solution until the red staining of the membrane disappear. 25. Start from step 11 using a second set of primary and secondary antibodies.

4

Notes 1. 4 Laemmli buffer: 0.5 M Tris–HCl pH 6.8, 8% (w/v) SDS, 40% (v/v) glycerol, 0.05% (w/v) Bromophenol blue.

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2. 2,2,2-Trichloroethanol is added to the acrylamide:bisacrylamide 30% (v/v) stock solution to visualize proteins that have migrated inside the gel by UV detection using the Stain-free procedure using a ChemiDoc Touch Imaging System. This procedure can serve as loading control to assess that all lanes were initially loaded with comparable amounts of proteins. Alternatively, Red Ponceau staining of the membrane can be performed after transfer to verify the migration and transfer. 3. In order to remove the parental plasmid DNA used as a matrix for PCR amplification, DpnI digestion can be performed. For this, the isothermal assembly reaction is supplemented with 2 μL of 10 FastDigest buffer and 10 U (1 μL) DpnI (Thermo Fisher) and incubated at 37  C for 1 h. Subsequently, DpnI is heat-inactivated (5 min at 95  C). Removing the template DNA from the isoT reaction decreases the possibility to transform E. coli cells with an empty plasmid and thus the number of false positives. 4. The cell suspension should be milky without being too dense. Usually for a volume of 1 mL of an overnight culture, 100 μL deionized water is used to resuspend the pellet. 5. Make sure the salmon sperm DNA is not too hot when adding the cells to avoid heat shock, which would cause cell death and thus reduce transformation efficiency. 6. The pellet should not be resuspended when deionized water is added after heat shock. Addition of water will dilute the transformation mix. 7. To avoid bubbles, even in the middle of the coverslip, press on the corners. Pressing in the middle of the coverslip would cause cell burst. 8. Ensure to remove all the droplets when discarding the protein precipitation solution. Residual trichloroacetic acid droplets will change the blue color of Laemmli solution to a yellow/ orange color. This indicates that pH of the sample is too acid which can cause troubles during SDS-PAGE electrophoresis. 9. If the solution has a yellow or orange color after adding the 1 Laemmli solution to the cells, Tris–Base 1 M should be added until the color turns to deep blue (start using 1 μL Tris–Base and increase the volume if the color does not change). Laemmli buffer 1 can also be supplemented with 1 M Tris–Base. 10. We used the Trans-Blot Turbo transfer system from BIO-RAD for our experiments, but any transfer system can be used. 11. The primary antibody can also be incubated at RT for 3 h under shaking. However, better results were obtained with an ON incubation at 4  C.

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12. Red Ponceau staining solution is used to denaturate the HRP coupled to the secondary antibody. By doing so, it is possible to incubate the membrane with another set of primary and secondary antibodies, without being disturbed by the HRP signal emitted by the first secondary HRP-coupled antibody. However, this method does not remove HRP-coupled secondary antibody from membrane. In order to reuse the membrane with a second set of antibodies, other stripping methods (heat and detergent, low pH or kits provided by different manufacturers) can be used.

Acknowledgments This work of the Interdisciplinary Thematic Institute IMCBio, as part of the ITI 2021-2028 program of the University of Strasbourg, CNRS and Inserm, was supported by IdEx Unistra (ANR10-IDEX-0002), and by SFRI-STRAT’US project (ANR 20SFRI-0012) and EUR IMCBio (ANR-17-EURE-0023) under the framework of the French Investments for the Future Program (to H.D.B, M.H, B.S); the National Science Center of Poland grant nr UMO-2018-31-B-NZ3-01117 (to R.K); and the Partenariat Hubert Curien Polonium 2021 Program Projet No. 44912NJ. References 1. Ben-Menachem R, Pines O (2017) Detection of dual targeting and dual function of mitochondrial proteins in yeast. Methods Mol Biol 1567:179–195 2. Ben-Menachem R, Tal M, Shadur T, Pines O (2011) A third of the yeast mitochondrial proteome is dual localized: a question of evolution. Proteomics 11:4468–4476 3. Kisslov I, Naamati A, Shakarchy N, Pines O (2014) Dual-targeted proteins tend to be more evolutionarily conserved. Mol Biol Evol 31:2770–2779 4. Lang A, John Peter AT, Kornmann B (2015) ER-mitochondria contact sites in yeast: beyond the myths of ERMES. Curr Opin Cell Biol 35: 7–12 5. Elbaz-Alon Y, Rosenfeld-Gur E, Shinder V, Futerman AH, Geiger T, Schuldiner M (2014) A dynamic interface between vacuoles and mitochondria in yeast. Dev Cell 30: 95–102 6. Cabantous S, Terwilliger TC, Waldo GS (2005) Protein tagging and detection with engineered self-assembling fragments of green fluorescent protein. Nat Biotechnol 23: 102–107

7. Bader G, Enkler L, Araiso Y, Hemmerle M, Binko K, Baranowska E, De Craene OJ, RuerLaventie J, Pieters J, Tribouillard-Tanvier D, Senger B, di Rago J-P, Friant S, Kucharczyk R, Becker HD (2020) Assigning mitochondrial localization of dual localized proteins using a yeast Bi-Genomic Mitochondrial-Split-GFP. elife 9:e56649 8. Bonnefoy N, Fox TD (2001) Genetic transformation of Saccharomyces cerevisiae mitochondria. Methods Cell Biol 65:381–396 9. Pedelacq JD, Cabantous S, Tran T, Terwilliger TC, Waldo GS (2006) Engineering and characterization of a superfolder green fluorescent protein. Nat Biotechnol 24:79–88 10. Inoue H, Nojima H, Okayama H (1990) High efficiency transformation of E. coli with plasmids. Gene 96:23–28 11. Frechin M, Senger B, Braye M, Kern D, Martin RP, Becker HD (2009) Yeast mitochondrial Gln-tRNA(Gln) is generated by a GatFABmediated transamidation pathway involving Arc1p-controlled subcellular sorting of cytosolic GluRS. Genes Dev 23:1119–1130

Chapter 17 Analysis of Mitochondrial Performance in Lymphocytes Using Fluorescent Lifetime Imaging Microscopy Meha Patel, Javier Manzella-Lapeira, and Munir Akkaya Abstract During lymphocyte maturation and differentiation, cells undergo a series of proliferative stages interrupted with stages of low activity. The rapid proliferation stages are marked by changes in metabolic outputs— adapting to energy demands by either hindering or utilizing metabolic pathways. As such, it is necessary to view these changes in real time; however, current strategies for metabolomics are time consuming and very rarely provide a holistic profile of the cellular metabolism while also characterizing mitochondrial metabolism. Here, we devised a fluorescence lifetime imaging microscopy (FLIM) strategy to image mitochondrial metabolic profiles in lymphocytes as they go through changes in metabolic activity. Our method provides not only a comprehensive view of cellular metabolism but also narrow in mitochondrial contributions while also efficiently excluding non-viable cells with and without the use of a viability dye. Our novel imaging strategy offers a reliable tool to study changes in mitochondrial metabolism. Key words Lymphocytes, Microscopy, FLIM, Immunometabolism, Immunology, Mitochondria

1

Introduction Lymphocytes are an integral part of mounting and maintaining immune response. To launch immune response, cells need to go through several processes with distinct metabolic states. These include activation, proliferation, and differentiation. Recently, focus on lymphocyte maturation has been centered around the metabolic changes that mark these stages [1, 2]. As such, it is important to be able to analyze the metabolic profile of lymphocytes which includes understanding their mitochondrial efficiency. The metabolic state of the cell can be evaluated using tests such as glucose and amino acid uptake, reactive oxygen species (ROS) production, real-time metabolic flux assays, as well as mitochondria-specific fluorescent reagents and antibodies used in conjunction with flow cytometry and microscopy [3–6].

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Recent imaging strategies such as confocal, stimulated emission depletion (STED), spinning disk, employ use of fluorescent markers—either endogenously or via antibodies—that target specific markers either on the surface or within mitochondria [6, 7]. However, most of these strategies have relied on the fluorescent markers to accurately pinpoint mitochondria in larger cell volumes than found in most immune cells [8]. Additionally, lymphocytes do not fare well in ex vivo cultures and quickly lose viability under experimental conditions [9]. Therefore, discriminating viable cells from dead cells as well as reducing the disruptions to mitochondria via staining is a critical factor that needs to be addressed within the capacity of metabolic assays. These difficulties, along with other hinderances such as the poor ability of lymphocytes to adhere to mounting surfaces, introduced the need to develop lymphocytespecific experimental strategies for visualizing mitochondrial metabolism. Here we provide a protocol for inspecting mitochondria metabolism in lymphocytes using fluorescence lifetime microscopy (FLIM), which measures the characteristic time that a molecule remains in the excited state prior to emitting a photon and returning to the ground state. FLIM is used to assess the abundance of metabolites, in particular oxidized and reduced forms of Nicotinamide adenine dinucleotide, abbreviated as NAD+ and NADH, respectively, and enzyme-bound NADH [10]. Given that these molecules autofluorescence, it obviates having to tag them with an endogenous fluorophore. When NADH is bound to enzymes, its autofluorescence lifetime increases compared to the free NADH molecule. Furthermore, when NADH is oxidized to NAD+, the molecule is no longer autofluorescent [11]. These properties elucidate the metabolic processes that are taking place in the mitochondria. In conventional confocal microscopy, there is a reliance on use of fluorochromes to outline or mark mitochondrial content, as well as other factors such as viability, glucose transporters, etc. [5, 11]. Under confocal microscopy, the accumulation of different fluorochromes may convolute signals. Therefore, we benefited greatly from the use of fluorescence lifetime microscopy in our protocol which not only provides a distinction between whole-cell and mitochondrial metabolic output but also allows us to minimize the use of fluorochromes [12, 13]. FLIM uses a combination of confocal and two-photon microscopy to excite bound and unbound NADH. Therefore, FLIM does not require fluorescent markers and in fact frees up laser channels for additional read outs to be combined with very little crossover. Following image acquisition, we process the image to subtract dead cells from live cells, while also distinctly outlining mitochondria using tetramethylrhodamine, methyl ester (TMRM) as a marker as well as an additional viability marker, LIVE/DEAD Far Red.

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In conclusion, our protocol provides mitochondrial metabolomics with the use of FLIM in order to eliminate conventional staining and imaging methods.

2

Materials

2.1 Isolation of Lymphocytes

1. MACS Buffer: Sterile PBS, supplemented with 0.5% BSA and 2 mM EDTA (degassed by filtering through a 0.22 μM filter). 2. 70 μM Cell strainer. 3. 30 μM Cell Strainer. 4. Magnetic cell isolation kit (e.g., Miltenyi Biosciences). 5. Miltenyi Magnetic Separators (e.g., Midimacs Magnetic Separator). 6. Biotin CD93 and CD9 antibodies (ThermoFisher). 7. Growth media: RPMI-1640 supplemented with 10% (v/v) heat-inactivated Fetal Bovine Serum, 1% penicillin–streptomycin, 2% glutamine, 1% HEPES, 1% Sodium Pyruvate, 1% MEM Non-Essential Amino Acids, and 0.1% 2-Mercaptoethanol (see Note 1).

2.2

Cell Staining

1. Staining buffer: Sterile PBS. 2. Viability stain: LIVE/DEAD Fixable Far Red Dead Cell Staining Kit (ThermoFisher): Prepare a stock solution by dissolving lyophilized dye in 50 μL sterile DMSO and store at 20  C. For working solution, prepare a 1:250 dilution of the stock solution in an appropriate volume of staining buffer immediately before use. 3. Tetramethylrhodamine, Methyl Ester, Perchlorate (TMRM) (ThermoFisher) staining solution: Prepare a stock solution by dissolving TMRM dye in 5 mL of sterile DMSO and store at 20  C. For a working solution, dilute 5 μL of stock in 1 mL of media as an intermediate dilution. Further dilute by adding 12 μL of intermediate dilution solution to 388 μL media immediately before use. TMRM staining is utilized to locate the mitochondria and limit our FLIM assay analysis to the mitochondria. While FLIM can be utilized for whole-cell metabolism here we use it to characterize mitochondrial metabolism. 4. RPMI 1640 without phenol red: RPMI 1640 without phenol red with 10% (v/v) heat-inactivated Fetal Bovine Serum, 1% penicillin–streptomycin, 2% glutamine, 1% HEPES, 1% Sodium Pyruvate, 1% MEM Non-Essential Amino Acids, and 0.1% 2-Mercaptoethanol (see Note 1).

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2.3 Slide Preparation and Mounting

1. Labtek eight-well chambered coverglass w/cvr #1 German borosilicate sterile (ThermoFisher Scientific, 155411). 2. 0.01% poly-L-lysine solution. 3. Slide preparation: Under sterile conditions (see Note 2), add 160 μL of 0.01% poly-L-lysine to the center of the middle coverslips or as many as needed to cover the bottom of the well entirely without excess coming up the walls. Incubate at room temperature for 5 min with the top off. After incubation, aspirate the poly-L-lysine and allow the coverslip to dry at room temperature for at least 2 h prior to plating and imaging (see Note 3).

2.4 Microscopy and Analysis

1. Cinefoil; matte black aluminum to mask light. 2. Zeiss LSM 780 with a 63 objective and an incubation system. 3. Chameleon Coherent laser. 4. Becker & Hickl TCSPC hardware (DCC-100 control card) with two HCM-100 detectors. 5. Becker & Hickl Spcm64 acquisition software and SPCImage analysis software. 6. MATLAB software (Mathworks).

3

Methods

3.1 Lymphocyte Isolation

1. This protocol is applicable for studying cell mitochondria as such it is applicable for most cells. For this protocol, we used follicular B lymphocytes freshly isolated from mouse spleens using negative magnetic isolation strategies described previously in detail [9, 14, 15] (see Note 4). 2. C57BL/6 (Stock no. 00664) was purchased from Jackson Laboratory and maintained at NIAID animal facilities according to Animal Care and Use Committee (ACUC) standards. Experiments involving mice were approved by NIH ACUC. 3. Seed conditions in a six-well plate at 6  106 B cells per well in 1.5 mL of RPMI 1640 media supplemented with 10% (v/v) heat-inactivated Fetal Bovine Serum, 1% penicillin–streptomycin, 2% glutamine, 1% HEPES, 1% Sodium Pyruvate, 1% MEM Non-Essential Amino Acids, and 0.1% 2-Mercaptoethanol and place in a 37  C incubator for 24 h (see Note 5) (Fig. 1).

3.2 Cell Staining in Solution: LIVE/DEAD Fixable Far Red Cell Stain and TMRM

1. Transfer cell solution to 1.5-mL centrifuge tubes and centrifuge at 600  g for 3 min at 4  C, then discard supernatant. Immediately wash well with 1 mL of sterile PBS and add to tubes then centrifuge at 600  g for 3 min at 4  C, then discard supernatant (see Notes 6 and 7).

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Fig. 1 Overview of the staining and imaging protocol

2. Gently resuspend cells in 100 μL LIVE/DEAD Fixable Far Red Cell Stain working solution at a maximum density of 4  106 B lymphocytes/mL (see Note 8). 3. Incubate cells on ice or at 4  C, protected from light for 20 min. After incubation, rinse cells with staining buffer, centrifuge as above, and discard supernatant.

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4. Gently resuspend cells in TMRM working solution at a density of 2  106 B lymphocytes/mL. 5. Incubate cells in a 37  C water bath for 30 min (see Note 9). 6. After incubation, rinse cells with RPMI 1640 media without phenol red supplemented with 10% (v/v) heat-inactivated Fetal Bovine Serum, 1% penicillin–streptomycin, 2% glutamine, 1% HEPES, 1% Sodium Pyruvate, 1% MEM Non-Essential Amino Acids, and 0.1% 2-Mercaptoethanol. Centrifuge as above and discard supernatant and resuspend in media. 7. Cells to be imaged should be resuspended in media at an optimal concentration depending on cell size and morphology determined by user; however, we suggest 2 million B lymphocytes/300 μL as a reference (see Note 10). 3.3 Adhesion of Cells to Slide

1. Carefully add 300 μL of the cell suspension to the center of a 0.01% poly-L-lysine treated chamber well. Incubate for 5 min in dark, 37  C incubator for 5 min (Fig. 1). 2. Fill each chamber well with up to 400 μL of RPMI 1640 without phenol red supplemented with 10% (v/v) heatinactivated Fetal Bovine Serum, 1% penicillin–streptomycin, 2% glutamine, 1% HEPES, 1% Sodium Pyruvate, 1% MEM Non-Essential Amino Acids, and 0.1% 2-Mercaptoethanol. The less media needed the better; therefore, 300 μL is more than enough and, if possible, utilize less.

3.4 Microscope Setup

This protocol uses a Coherent Chameleon II Ti:Sapphire femtosecond pulsed infrared laser coupled with a Zeiss LSM 780 confocal microscope. The FLIM acquisition is initiated with the Zen software, while a Becker & Hickl TCSPC DCC-100 control card with two HCM-100 detectors and the Spcm64 software acquire the photon data into .sdt files. This protocol can be adapted to any confocal/FLIM system. 1. Assemble stage heating devices and attach the lens heater to the Plan-Apochromat 63 1.4 NA oil-immersion objective lens. Set the temperature to 37  C at least 1 h before imaging. 2. Turn on the multiphoton laser and the FLIM detector. 3. On the FLIM computer, open Spcm64. With two detectors, place a dichroic separating the emission at 525 nm, with a BFP filter (480/40) for the reflected shorter wavelength light (see Note 11). 4. Open the System Parameters for each detector. Operation mode should be set to FIFO. Set collection, display, and repeat times to 1 s. Set ADC resolution to 256. The “Image Pixels X” and “Image Pixels Y” should match the pixel resolution that

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you define with the Zen configuration. Save and close this window. 5. On the main panel, match the “SYNC” bar matches to the laser repetition rate of 80 MHz. It should read 8.00E+7. 6. Open the ZEN software and configure the parameters for the acquisition of a DIC/fluorescence image and for the FLIM scanning. 7. For the confocal configuration, select “LSM” and “Channel” mode, with line switching. 8. Track 1: activate the 561 nm laser then select the GaAsP detector to collect for the signal between 570 and 630 nm. Select a beam-splitter that includes both 561 and 633. Check the T-PMT box to acquire a DIC image. 9. Track 2: activate the 633 nm laser then select the far-red sensitive PMT to collect signal between 640 and 730 nm. 10. Set the image size to 512  512 pixels, at 2 zoom with a 16-bit depth. Set the scan speed to 9 and average two times. Save the configuration. 11. For the FLIM configuration, select “Non-Descanned” and “Channel” mode. Activate the multiphoton laser and tune to 750 nm. Select the 690+ reflector off the invisible light path and select the MP 355/690 beam splitter. Make sure image size in pixels matches the system parameters on your FLIM detectors. Save this second configuration. 3.5 Image Acquisition

1. In order to minimize any light source while the FLIM detectors are activated, use the cinefoil to cover any indicator lights and LED screens on peripheral devices. The computer screens should be facing away from the microscope and dimmed to at least half their brightest level. 2. Place the eight-well chamber with the prepared cells on the stage and wait at least 10 min for the temperature to reach 37  C. 3. With the Zen software in the confocal configuration that was saved in Subheading 3.4, step 10, find the right focus and field of view. 4. With “Range indicator” checked, adjust gain and offset settings for the PMTs, as well as laser powers for the TMRM and Live/ Dead Far-Red markers. 5. Click “Snap” then click “New” and switch to the FLIM configuration saved in Subheading 3.4, step 11. 6. In Spcm64, enable the FLIM detectors in the DCC-100 control window. There should be some level of background photons detected, but the highest value on the CFD should

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Fig. 2 Demonstration of viable and non-viable cells. Freshly isolated follicular B lymphocytes were cultured for 24 h in media alone (left) or media supplemented with Anti-IgM antibody that triggers B-cell receptor signaling (right). Following culture, cells were stained with LIVE/DEAD (red) and TMRM (yellow) and imaged using confocal microscopy. In both the cases, TMRM has been shown to stain only viable cells that are negative for the LIVE/DEAD stain

not be greater than 102. If it is 103, you need to check for background light sources and make sure you cover them. 7. Click “Start” on the menu. The windows should be white, waiting for the trigger from the scanner. On Zen, click “Continuous” acquisition. The FLIM detectors will begin collecting events and the image will be updated every second with new photons. CFD should be in the 103 to 105 range (see Note 12). 8. Aim to collect 20–30 million events, which is approximately 60 s of continuous scanning. 9. Click “Stop” on the menu, then stop the acquisition on Zen. Save the FLIM file. 10. Return to the confocal configuration in Zen and repeat steps 3–9, making sure to acquire at least 30 cells per experimental condition for statistical analysis (Fig. 2). 3.6

FLIM Analysis

1. Open the SPCImage software and select “Import” from the “File” menu to load the data for a raw FLIM data file. 2. Select double-exponential photon decay model: Under “Multiexponential Decay” on the right side of the screen, select two components. Set the bin to 3 and the threshold to 8. 3. Open the model options window and specify the maximum Chi-squared value as 2.0 (see Note 13).

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Fig. 3 Comparison of different metabolic activation within color-coded FLIM images. Freshly isolated follicular B lymphocytes were stained as described in Fig. 2, after being cultured for 24 h in media (left) or media supplemented with Anti-IgM stimulated (right). After confirming the cellular viability using confocal as seen in Fig. 2, images are then subject to data analysis via SPCImage software to render this color-coded FLIM signal image. The colors indicate the lifetime FLIM signals with our lower range at 26–1638 ps. Unbound NAD+ tends to have shorter lifetimes while bound NADH has longer lifetimes [10]

4. To change the image properties, right click on the image and select “Intensity.” This will open a window that allows you to change the intensity and contrast for both the grayscale fluorescence and the color-coded FLIM signal. Right click on the colored histogram to the left of the image. Select “Color,” which will open a window where you can change the range values of the color-coded image (Fig. 3). 5. Select “Decay Matrix Single Channel” under the “Calculate” menu. After processing, click on “Phasor Plot” to create the Phasor components for each pixel. The phasor will be displayed in a new window. 6. After you have processed the initial file, you can select the batch processing option under the “Calculate” menu for that channel. I will provide a file selector menu in which multiple files can be selected for batch processing. 7. In order to obtain the data matrices for each image, select the “Export” option under the “File” menu. 8. Repeat this data processing for each experimental group. A custom-made MATLAB algorithm can then be used to exclude the cells that have the cell-death fluorescence marker and to

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Fig. 4 Demonstration of the staining and image analysis strategy in B lymphocytes that shows two distinctive populations: unstimulated (yellow) and Anti-IgM stimulated (green). Follicular B Lymphocytes were stained with LIVE/DEAD to exclude dead cells and TMRM to define mitochondrial boundaries and imaged using confocal and FLIM microscope settings. The representative phasor plot showing mitochondrial performance in individual cells (small dots) and the mean of each condition (large dots) are shown

segment the mitochondrial regions in order to produce the final phasor plots (Fig. 4).

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Notes 1. For this method, the media prepared was utilized for ideal B lymphocyte conditions. The media for B lymphocyte conditions was prepared utilizing 500 mL of RPMI 1640 with and without phenol red, supplemented with 50 mL heatinactivated fetal bovine serum, 12 mL Pen-Strep/L-Glut, 6 mL HEPES, 6 mL Sodium Pyruvate, 6 mL MEM Non-Essential Amino Acids, and 600 μL 2-mercaptoethanol. However, given the versatility of this method, utilizing media supplemented to support the chosen cell type is ideal. 2. It is not absolutely necessary to maintain a sterile environment for the purposes of cell preparation, it is helpful to carry out the coverslip preparation in a biosafety cabinet to avoid introduction of dirt or dust onto the well. You may find it helpful to label the sides of the well near the mouth with permanent

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marker as the bottom will come into contact with oil immersion fluid and the lid will be taken off for acquisition purposes. 3. The coated coverslip may be prepared and dried overnight, if kept free of contaminants such as dust. 4. Our B lymphocyte isolation strategies utilize magnetic separation which allows for no antibody-labeling during isolation. For the sensitivity of this imaging technique, utilizing no to very little antibody labeling will likely reduce the amount of interference seen under acquisition. 5. For the sake of this protocol, we seeded unstimulated vs. stimulated as a rope comparison. On a normal coverslip, you may seed up to four conditions. 6. The cells stained with LIVE/DEAD and/or TMRM may be stored at 4  C for up to 30 min after thorough washing with staining buffer and resuspension in media. In order to maintain cell viability, the time spent stored should be minimized. 7. From this point till step 6, the cells should remain in staining buffer. 8. Optional: If you do not want to stain cells with LIVE/DEAD, omit steps 2 and 3. 9. Incubation of the cells can take place either in a water bath at 37  C for 30 min or within an incubator at 37  C and 5% CO2. Be sure to check temperatures as TMRM stains ideally at 37  C. 10. While we suggest 2 million cells/300 μL, this will need to be titrated to fit the user’s preferences. In most cases, keeping seeding count at an optimal level, using as little media as possible is suggested. The minimum amount of media used should be 160 μL so as to coat the bottom of the coverslip entirely. This allows for less interference with image acquisition. 11. This is the autofluorescence FLIM channel for various components including hemoglobin and conjugated/unconjugated NAD(P)H. If the user has only one FLIM detector, it should be used with a CFP filter for the NAD(P)H channel. 12. If the CFD value is above this range, the laser power is too high and might damage the cells. Laser power recommended is 1.5 mW at the objective. The user might want to test various powers. 13. The chi-squared value measures how accurate the data is fit to the mathematical model. The user can threshold the data based on the goodness of the fit, which is indicated by how much the chi-squared value differs from a value of 1.

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Acknowledgments This work was supported by the Intramural Research Program of the National Institutes of Health, National Institute of Allergy and Infectious Diseases. Dr. Munir Akkaya is funded by the Ohio State University College of Medicine. The authors thank Ryan Kissinger for drawing the illustration used in Fig. 1 and Joe Brzostowski for his valuable input to this protocol. References 1. Akkaya M, Pierce SK (2019) From zero to sixty and back to zero again: the metabolic life of B cells. Curr Opin Immunol 57:1–7. https://doi. org/10.1016/j.coi.2018.09.019 2. O’Neill LAJ, Kishton RJ, Rathmell J (2016) A guide to immunometabolism for immunologists. Nat Rev Immunol 16(9):553–565. https://doi.org/10.1038/nri.2016.70 3. Kurupati RK, Haut LH, Schmader KE, Ertl HCJ (2019) Age-related changes in B cell metabolism. Aging (Albany NY) 11(13): 4367–4381. https://doi.org/10.18632/ aging.102058 4. Russell S, Wojtkowiak J, Neilson A, Gillies RJ (2017) Metabolic profiling of healthy and cancerous tissues in 2D and 3D. Sci Rep 7(1): 1–11. https://doi.org/10.1038/s41598-01715325-5 5. Akkaya M, Traba J, Roesler AS et al (2018) Second signals rescue B cells from activationinduced mitochondrial dysfunction and death. Nat Immunol 19(8):871–884. https://doi. org/10.1038/s41590-018-0156-5 6. Cottet-Rousselle C, Ronot X, Leverve X, Mayol JF (2011) Cytometric assessment of mitochondria using fluorescent probes. Cytom Part A 79A(6):405–425. https://doi. org/10.1002/cyto.a.21061 7. Godet I, Shin YJ, Ju JA, Ye IC, Wang G, Gilkes DM (2019) Fate-mapping post-hypoxic tumor cells reveals a ROS-resistant phenotype that promotes metastasis. Nat Commun 10(1): 1–18. https://doi.org/10.1038/s41467019-12412-1 8. Chan YK, Tsai MH, Huang DC, Zheng ZH, Hung KD (2010) Leukocyte nucleus segmentation and nucleus lobe counting. BMC Bioinformatics 11:558–558. https://doi.org/10. 1186/1471-2105-11-558

9. Traba J, Miozzo P, Akkaya B, Pierce SK, Akkaya M (2016) An optimized protocol to analyze glycolysis and mitochondrial respiration in lymphocytes. J Vis Exp 2016(117): 54918. https://doi.org/10.3791/54918 10. Blacker TS, Duchen MR (2016) Investigating mitochondrial redox state using NADH and NADPH autofluorescence. Free Radic Biol Med 100:53–65. https://doi.org/10.1016/j. freeradbiomed.2016.08.010 11. Blacker TS, Mann ZF, Gale JE et al (2014) Separating NADH and NADPH fluorescence in live cells and tissues using FLIM. Nat Commun 5(1):3936. https://doi.org/10.1038/ ncomms4936 12. Sekar RB, Periasamy A (2003) Fluorescence resonance energy transfer (FRET) microscopy imaging of live cell protein localizations. J Cell Biol 160(5):629–633. https://doi.org/10. 1083/jcb.200210140 13. Jabbour JM, Cheng S, Malik BH et al (2013) Fluorescence lifetime imaging and reflectance confocal microscopy for multiscale imaging of oral precancer. J Biomed Opt 18(04):1. https://doi.org/10.1117/1.JBO.18.4. 046012 14. Akkaya B, Holstein AH, Isaac C et al (2017) Ex-vivo iTreg differentiation revisited: convenient alternatives to existing strategies. J Immunol Methods 441:67–71. https://doi. org/10.1016/j.jim.2016.11.013 15. Akkaya B, Miozzo P, Holstein AH, Shevach EM, Pierce SK, Akkaya M (2016) A simple, versatile antibody-based barcoding method for flow cytometry. J Immunol 197(5): 2027–2038. https://doi.org/10.4049/ jimmunol.1600727

Chapter 18 Monitoring Mitochondrial Perturbations During Infection Varnesh Tiku and Man-Wah Tan Abstract Mitochondria are pivotal organelles in the cell that regulate a myriad of cellular functions, which eventually govern cellular physiology and homeostasis. Intriguingly, microbial infection is known to trigger morphological and functional alterations of mitochondria. In fact, a number of bacteria and viruses have been reported to hijack mitochondrial functions including cell death induction and regulation of immune signaling to evade detection, promote their intracellular growth and subsequent dissemination. Here we describe methodologies that can be applied to assess mitochondrial functions upon infection. More specifically, we outline experimental procedures used to evaluate different parameters including mitochondrial morphology, adenosine triphosphate (ATP) levels, reactive oxygen species (ROS) levels, and mitophagy. Together these parameters can help gauge the overall health of mitochondria upon infection. Key words Mitochondria, Bacterial and viral infection, Mitochondrial fission and fusion, ROS levels, ATP levels, Mitophagy

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Introduction Mitochondria regulate a variety of cellular functions that impact cellular physiology. They are very dynamic organelles that undergo cycles of fission and fusion giving rise to elongated tubular networks or shorter fragmented structures [1, 2]. These morphological states govern mitochondrial function. Mitochondrial fission and fusion are modulated by the GTPase proteins Mitofusin 1 and 2 (MFN1 and MFN2), optic atrophy 1 (OPA1), and dynamin related protein 1 (DRP1) [3]. During mitochondrial fusion, shorter fragments of mitochondria are clipped together to form elongated networks. Outer mitochondrial membrane proteins MFN1 and MFN2 facilitate the joining of outer mitochondrial membranes while the inner membrane protein OPA1 induces the fusion of the inner mitochondrial membrane [2, 4] (Fig. 1a). In contrast, conditions that activate mitochondrial fission enhance the accumulation of DRP1 on mitochondria, which induces scission of longer tubules into shorter mitochondrial fragments [5] (Fig. 1a).

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Fig. 1 Mitochondrial fission and fusion pathways. (a) The illustration depicts mitochondrial proteins involved in the fission and fusion pathways (created with BioRender.com) (b) A549 lung epithelial cells infected with the Gram-negative bacterial pathogen A. baumannii at a multiplicity of infection (MOI) of 50 for 6 h. Immunofluorescence was performed following the procedure detailed in the Subheading 3.1. Mitochondria are stained with anti-TOM20 antibody (red), and the nuclei are labeled with DAPI (blue). Mitochondrial fragmentation is clearly visible in the infected cell compared to the uninfected cell. Scale bar represents 10 μm

Elongated tubular mitochondria display higher levels of oxidative phosphorylation (OXPHOS) and thus generate high levels of ATP. Electron leakage from the electron transport chain complexes in the mitochondria can lead to partial reduction of oxygen to form superoxides which collectively are termed reactive oxygen species (ROS). Fragmented mitochondria are associated with lower levels of ATP production and higher build-up of ROS [6]. Given its

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pivotal function in cells, it is not surprising that a variety of microorganisms have evolved mechanisms to hijack the host mitochondrial functions to promote infectivity and survival [2, 7]. As a consequence, bacterial and viral infections can often perturb mitochondrial dynamics, thereby affecting its morphology and eventually impacting its cellular functions [2, 7–10] (Fig. 1b). To maintain cellular homeostasis, mitochondrial fitness is closely monitored in cells. As such, damaged mitochondria, due to infection or other insults, are rapidly cleared via a dedicated process known as mitophagy. Mitophagy is mediated via the action of a mitochondrial serine/threonine kinase PINK1 and one of its substrates, Parkin that functions as an E3 ubiquitin ligase. Under physiological conditions, PINK1 is degraded by mitochondrial proteases [11]. However, upon mitochondrial damage, mitophagy is triggered through the stabilization of PINK1 on the mitochondrial membrane and the subsequent phosphorylation of ubiquitin and Parkin, which activates Parkin [12] (Fig. 2). This is followed by an accumulation of activated Parkin on mitochondrial membrane that leads to the assembly of mono- and poly-ubiquitin chains that serve as signals for the recruitment of the autophagy machinery, thereby targeting damaged mitochondria for lysosomal degradation (Fig. 2). In particular, damaged mitochondria harboring the ubiquitin signal are engulfed by specialized doubled membrane vesicles known as autophagosomes that then fuse with lysosomes, delivering the damaged mitochondria in the process [12] (Fig. 2). As such, a clear indicator of mitophagy is the co-localization of the autophagosomal protein LC3 with mitochondria. Bacterial and viral infections are known to impact host cell mitophagy [13, 14]. Restoration of organelle homeostasis through mitophagy is an attempt by the cell to remove damaged mitochondria and reinstate normal cellular physiology. However, if mitochondria become highly fragmented and impaired beyond repair during infection, this eventually triggers cell death [2, 15]. Therefore, probing mitochondrial morphology, assessing the levels of ATP and ROS and examining mitophagy induction serve as powerful tools to interrogate mitochondrial function upon infection. In the following sections, we will describe in detail the tools and methodologies that are utilized and well accepted in the field to assess mitochondrial morphology, ROS, and ATP levels as well as mitophagy. Mitochondrial morphology can be visualized via performing immunofluorescence using mitochondrial marker proteins like TOM20 followed by imaging with a confocal microscope using high magnifications. ROS levels can be detected using fluorogenic probes, which when oxidized inside the cells, emit strong fluorescence. The function of mitochondria can be assessed via the quantification of ATP levels using luminescent ATP substrates. Additionally, mitochondrial damage can be evaluated by examining

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Fig. 2 The mitophagy pathway. The figure is an illustration of the mitophagy pathway. During mitophagy, PINK1 and Parkin accumulate on the mitochondria. Parkin ubiquitinates multiple mitochondrial proteins which then serve as signals for the recruitment of the autophagy machinery for subsequent clearance of the damaged mitochondria. The autophagosomal membrane protein LC3 also accumulates on the damaged mitochondria. (Created with BioRender.com)

mitophagy induction using immunofluorescence. Below, each method is outlined in more detail.

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Materials

2.1 Visualization of Mitochondrial Morphology

1. 4% Paraformaldehyde (PFA) solution: Dilute 1 mL of 16% commercially available PFA solution in 3 mL of PhosphateBuffered Saline (PBS). 2. 0.1% Triton X-100: Add 100 μL of 10% Triton X-100 in 10 mL of PBS. 3. 1% Bovine Serum Albumin (BSA): Dissolve 0.1 g of BSA powder in 10 mL PBS. 4. Commercially available anti-TOM20 antibody and a corresponding fluorescently conjugated secondary antibody. 5. Commercially available mounting medium containing a DNA stain like 40 ,6-diamidino-2-phenylindole (DAPI). 6. Tissue culture treated four-well chamber slides. 7. A confocal microscope with 100 magnification.

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2.2 Examination of Mitochondrial Function

1. Commercially available fluorogenic probes for ROS detection.

2.2.1 ROS Assay

3. 30% hydrogen peroxide (H2O2) in sterile water.

2. 4% Paraformaldehyde (PFA) solution: Dilute 1 mL of 16% commercially available PFA solution in 3 mL of PBS. 4. Tissue culture treated four-well chamber slides. 5. Commercially available mounting medium containing a DNA stain like 40 ,6-diamidino-2-phenylindole (DAPI). 6. A confocal microscope with 100 magnification.

2.2.2 ATP Assay

1. Luminescent ATP binding probes. 2. White-walled clear bottom 96-well plate. 3. A plate reader with luminescence detection capability.

2.3 Visualization of Mitophagy

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For examining mitophagy, immunofluorescence is performed to visualize the mitophagy proteins PINK1 and Parkin and the autophagosomal marker LC3. The reagents listed in Subheading 2.1 are also required for examining mitophagy. In addition, commercially available anti-LC3, anti-PINK1, and anti-Parkin antibodies and corresponding fluorescently conjugated secondary antibodies are required. Researchers also use mitotracker, which is a fluorescent dye that labels mitochondria. Mitotracker can be used directly on live cells, thereby eliminating the need of the fixation step during immunofluorescence (see Note 4).

Methods

3.1 Analysis and Quantification of Mitochondrial Fission and Fusion

1. Seed the desired cells at a density of 105 cells per chamber in a four-chamber slide and perform the infection using the desired bacteria at the required multiplicity of infection (MOI) (see Note 1). 2. Following infection, wash the cells three times with PBS to remove extracellular bacteria. 3. Fix the cells with 4% PFA for 15 min at room temperature (see Notes 2 and 3). 4. Wash the fixed cells three times with PBS to remove PFA. Permeabilize the fixed cells with 0.1% Triton X-100 for 15 min at room temperature (see Note 4). 5. After permeabilization, wash the cells twice with PBS and block with 1% BSA for 1 h at room temperature. 6. After blocking, treat the cells with the anti-TOM20 antibody raised in rabbit or mouse. The antibody dilution should be performed in 1% BSA. Add 10 μL of anti-TOM20 antibody in 2 mL of 1% BSA to get a final dilution of 1:200. Add 400 μL

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of the diluted primary antibody to each chamber and incubate for 1 h at room temperature (see Note 5). 7. Wash the cells with PBS three times and treat the cells with the appropriate secondary antibody (mouse or rabbit) tagged with a fluorophore. The antibody dilution should be performed in 1% BSA. Add 4 μL of the secondary antibody in 2 mL of 1% BSA to get a final dilution of 1:500. Add 400 μL of the diluted secondary antibody to each chamber and incubate for 1 h at room temperature in the dark. 8. Wash three times with PBS and then remove all the liquid from all the chambers. Break open the walls of the chamber slide using the dedicated plastic tool (see Note 6). Dry out all the chambers by gently absorbing the leftover liquid using a tissue paper around the corners of the chambers. 9. Add a drop of mounting medium containing DAPI in the center of each chamber and gently drop a rectangular cover slip on top of the slide that covers the entire slide. 10. Keep the slide at room temperature in the dark overnight for the mounting medium to properly dry out and hold the coverslip onto the slide. 11. The slide can be imaged using a confocal microscope the next day. Typically, a laser scanning confocal microscope or a spinning disk confocal microscope can be used to image mitochondria and visualize differences in mitochondrial morphology. In certain cases, super resolution microscopy can be used to resolve minor differences in mitochondrial networks. 12. Typically, 15–20 images spanning different regions on the slide should be taken for the quantification of mitochondrial area and perimeter. 13. Differences in mitochondrial morphology can be easily assessed visually. However, it can be daunting to perform quantification of the differences in mitochondrial morphology. An open-source software called CellProfiler can be used to perform automated quantification of the mitochondrial area and perimeter [7, 16]. 14. In the CellProfiler program, after uploading the images, the program will automatically detect objects (DAPI-stained nuclei and TOM20-stained mitochondria). The objects touching the image borders should be eliminated to avoid artifacts in the quantification. 15. DAPI-stained nuclei should serve as the primary objects and the program will determine cell boundaries, which will enable the program to distinguish between individual cells and label the cells as secondary objects using the nuclei as the primary (also called the seed object in the program). Then the labeled

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mitochondria will be identified by the program through image segmentation using the watershed module of the program. Finally, the program will quantify the area, perimeter, and other shape features of mitochondria. The program then associates “child objects” (mitochondria) with the “parent objects” (cells) and determines the number of mitochondria associated with each cell and calculates mean measurement values for all the mitochondria that are associated with each individual cell. 16. The program exports all the measurement values to a spreadsheet from which the values can be plotted into graphs for comparisons and statistics. 17. The lower the mitochondrial area and perimeter, the higher the number of fragmented mitochondria and mitochondrial fission rate and vice versa. 3.2 Assessing ROS and ATP Levels 3.2.1 Examination of ROS Levels

1. Seed the desired cells at a density of 105 cells per chamber in a four-chamber slide and perform the infection using the desired bacteria at the required MOI. 2. For examining ROS levels, H2O2 treatment of cells can be used as a positive control. Cells can be treated with 30% H2O2 at a final dilution of 1:1000 for 1 h at 37  C. 3. After the infection or H2O2 treatment, add the fluorogenic probe at a final concentration of 5 μM and incubate at 37  C for 30 min (see Note 7). 4. Remove the medium and wash the cells three times with PBS. 5. Fix the cells with 4% PFA for 15 min at room temperature. 6. Remove the 4% PFA fixative and wash the cells three times with PBS. 7. Add a drop of mounting medium containing DAPI in the center of each chamber and gently drop a rectangular cover slip on top of the slide that covers the entire slide. 8. Image the cells with a confocal microscope within 24 h of mounting (see Note 8). 9. Cells displaying higher levels of ROS will exhibit stronger fluorescence. Fluorescence intensity can be quantified using standard open source image analysis programs such as Fiji (Image J).

3.2.2 Evaluation of ATP Levels

1. Seed the desired cells at a density of 105 cells per well in a white-walled clear bottom 96-well plate and perform the infection using the desired bacteria at the required MOI (see Note 9).

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2. After the infection, add 50 μL of commercially available cell lysis solution to each well and shake the plate for 5 min at 700 rpm in an orbital shaker. 3. Add 50 μL of luminescent ATP substrate solution to each well and shake the plate for 5 min at 700 rpm in an orbital shaker. 4. Incubate the plate for 10 min in the dark. 5. Measure luminescence in a plate reader to assess the levels of ATP. The higher the luminescence in a well, the higher are the ATP levels (see Note 10). 3.3 Evaluation of Cellular Mitophagy

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Mitophagy can be detected by examining the co-localization of PINK1 and Parkin with mitochondria. Additionally, upon the induction of mitophagy, the autophagosomal marker protein LC3 is also targeted to mitochondria. Therefore, examining the co-localization of PINK1, Parkin, and LC3 with mitochondria is a very well-accepted method of investigating the induction of mitophagy as illustrated in Fig. 2. Immunofluorescence is the most common method for tracking the localization of these proteins (see steps 1–11 in Subheading 3.1). The endogenous levels of PINK1 and Parkin are generally very low which makes detection via immunofluorescence hard. Therefore, many studies use cell lines overexpressing fluorescently tagged PINK1 and Parkin and monitor their co-localization with the mitochondrial protein TOM20.

Notes 1. It is easier to look at mitochondrial morphology in epithelial cells such as A549 and HeLa cells compared to immune cells like macrophages. Epithelial cells have a flattened morphology and are larger in size making it easier to detect morphological changes in mitochondria. 2. It is recommended to use PFA solution instead of methanol for fixation of cells because methanol fixation can lead to dehydration of the cells, which can shrink and distort membranous organelles like mitochondria. 3. After fixation with PFA solution, the cells can be washed three times with PBS and left at 4  C overnight to continue the rest of the procedure the next day. 4. While PFA fixation does permeabilize the cells to a certain degree, it is recommended to use 0.1% Triton X-100 for full permeabilization of the cells that will facilitate access of primary and secondary antibodies to intracellular targets for labeling mitochondria. Fixation and permeabilization are not be

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required if mitochondria labeling stains like mitotracker are used. 5. Primary antibody incubation can be performed overnight at 4  C. 6. The plastic tool that is used to remove the walls of the chamber slides comes with the slides, and there are clear instructions in the user manual on how to use the tool properly. After removing the chamber walls, make sure to remove the leftover glue that holds the walls together. The leftover glue can obstruct the coverslips from sticking to the slides which leads to trapping of air bubbles. 7. There is no need for removing the cell culture medium and washing the cells before adding the fluorogenic probe. It can be directly added into the cell culture medium. 8. Sometimes there is high background in the images, it is recommended to look at multiple different regions on the slide to assess lower background regions. It might also be worthwhile to perform this assay using two or three technical replicates to ameliorate the background issue. 9. Ensure that the final volume of the cell culture medium per well is 100 μL. This will be necessary since there will be further additions of the lysis solution and the ATP substrate solution into each well. 10. Ensure that the cell number is the same across different conditions. Differences in cell number can result in differences in the readings for ATP levels. References 1. Chan DC (2012) Fusion and fission: interlinked processes critical for mitochondrial health. Annu Rev Genet 46:265. https://doi. org/10.1146/annurev-genet110410-132529 2. Tiku V, Tan M-W, Dikic I (2020) Mitochondrial functions in infection and immunity trends in cell biology. Trends Cell Biol 30: 263. https://doi.org/10.1016/j.tcb.2020. 01.006 3. Sprenger H-G, Langer T (2019) The good and the bad of mitochondrial breakups. Trends Cell Biol 29:888. https://doi.org/10.1016/J. TCB.2019.08.003 4. Anand R, Wai T, Baker MJ et al (2014) The i-AAA protease YME1L and OMA1 cleave OPA1 to balance mitochondrial fusion and fission. J Cell Biol 204:919. https://doi.org/10. 1083/jcb.201308006

5. Pagliuso A, Cossart P, Stavru F (2018) The ever-growing complexity of the mitochondrial fission machinery. Cell Mol Life Sci 75: 355–374. https://doi.org/10.1007/s00018017-2603-0 6. Longo DL, Archer SL (2013) Mitochondrial dynamics-mitochondrial fission and fusion in human diseases. N Engl J Med 369: 2236–2251. https://doi.org/10.1056/ NEJMra1215233 7. Tiku V, Kofoed EM, Yan D et al (2021) Outer membrane vesicles containing OmpA induce mitochondrial fragmentation to promote pathogenesis of Acinetobacter baumannii. Sci Rep 11:618. https://doi.org/10.1038/s41598020-79966-9 8. Escoll P, Song O-R, Viana F et al (2017) Legionella pneumophila modulates mitochondrial dynamics to trigger metabolic repurposing of infected macrophages. Cell Host

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Microbe 22:302–316.e7. https://doi.org/10. 1016/J.CHOM.2017.07.020 9. Stavru F, Palmer AE, Wang C et al (2013) Atypical mitochondrial fission upon bacterial infection. Proc Natl Acad Sci U S A 110: 16003–16008. https://doi.org/10.1073/ pnas.1315784110 10. Jain P, Luo Z-Q, Blanke SR (2011) Helicobacter pylori vacuolating cytotoxin A (VacA) engages the mitochondrial fission machinery to induce host cell death. Proc Natl Acad Sci U S A 108:16032–16037. https://doi.org/ 10.1073/pnas.1105175108 11. Pickles S, Vigie´ P, Youle RJ (2018) Mitophagy and quality control mechanisms in mitochondrial maintenance. Curr Biol 28:R170 12. Palikaras K, Lionaki E, Tavernarakis N (2018) Mechanisms of mitophagy in cellular

homeostasis, physiology and pathology. Nat Cell Biol 20:1013 13. Zhang L, Qin Y, Chen M (2018) Viral strategies for triggering and manipulating mitophagy. Autophagy 14:1665 14. Zhang Y, Yao Y, Qiu X et al (2019) Listeria hijacks host mitophagy through a novel mitophagy receptor to evade killing. Nat Immunol 20:433–446. https://doi.org/10.1038/ s41590-019-0324-2 15. Xie LL, Shi F, Tan Z et al (2018) Mitochondrial network structure homeostasis and cell death. Cancer Sci 109:3686 16. McQuin C, Goodman A, Chernyshev V et al (2018) CellProfiler 3.0: next-generation image processing for biology. PLoS Biol 16: e2005970. https://doi.org/10.1371/journal. pbio.2005970

Chapter 19 Measuring the Mitochondrial Ubiquinone (Q) Pool Redox State in Isolated Respiring Mitochondria Marten Szibor, Estelle Heyne, Carlo Viscomi, and Anthony L. Moore Abstract The ubiquinone (Q) pool represents a node in the mitochondrial electron transport chain (ETC) onto which the electrons of all respiratory dehydrogenases converge. The redox state of the Q pool correlates closely with the electron flux through the ETC and is thus a parameter of great metabolic value for both the mitochondrial and cellular metabolism. Here, we describe the simultaneous measurement of respiratory rates of isolated mouse heart mitochondria and the redox state of their Q pool using a custom-made combination of a Clark-type oxygen electrode and a Q electrode. Key words Mitochondria, Redox state, Respiratory rates, Ubiquinone pool

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Introduction The mitochondrial electron transport chain (ETC) consists of four multi-subunit protein complexes and their integral redox partners, i.e., the ubiquinone (Q) pool and cytochrome c. In a series of steps, electrons are transferred through the ETC from specific substrates (NADH or succinate) to redox partners of increasing affinity with molecular oxygen being the ultimate acceptor. This electron transfer is coupled to proton pumping across the inner mitochondrial membrane, thereby generating an electrochemical gradient, which itself is the major driving force (protonmotive force, pmf) for ATP production at complex V (or F1F0 ATPase). Notably, all respiratory chain dehydrogenases such as NADH:ubiquinone oxidoreductase (complex I) and succinate dehydrogenase (complex II) facilitate substrate oxidation and Q pool reduction (Q reducing enzymes) while cytochrome bc1 complex (complex III) in conjunction with cytochrome c and cytochrome c oxidase (complex IV) or an alternative oxidase (AOX) facilitates Q pool oxidation (Q oxidizing enzymes) [1–3]. From this, it follows that the mitochondrial Q pool represents a functional node (bottle neck) [2, 4, 5] in electron

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flux through the ETC and hence renders it a parameter of great metabolic significance. Importantly, although mitochondrial respiration and the Q pool redox state are functionally interrelated, there is no linear relationship between the two [2]. Therefore, respiratory rates and the Q reduction state need to be measured separately. We have previously estimated both parameters simultaneously in isolated mitochondria by the combination of custommade Clark-type and Q electrodes [2, 4]. Technical details of such electrodes, including recently developed commercial devices, have previously been described in detail ([2, 4] and Dr P.R. Rich, UCL London, European Patent no. 85900699.1/). Briefly, oxygen levels were assessed using a Clark-type electrode. In this system, oxygen is reduced at the cathode and silver is oxidized at the anode. To prevent contamination, the platinum surface is covered with an oxygen-permeable membrane. Following a device-specific equilibration time, the resultant current thus reflects the rate of reaction and is critically determined by the diffusion of oxygen. The steady-state redox poise of the mitochondrial Q-pool can also be measured voltametrically as originally demonstrated by Dr P.R. Rich (UCL London, European Patent no. 85900699.1/). In this system, added Q-1 or Q-2 acts as a mimetic of the steady-state reduction level of the endogenous Q-pool by reacting with both Q reduction sites of the dehydrogenases and the Q-oxidation sites of the oxidases and the measuring electrode but not directly with the endogenous Q within the membrane. On the basis that the redox state of the Q mimetic is in equilibrium with the redox state of Q-pool in the respiratory chain via its interaction with the dehydrogenases and oxidases, the redox state of the Q mimetic reflects the redox state of the mitochondrial Q-pool [2, 4]. Indeed the Q-measurement system described below accurately reflects the redox levels of the Q-pool measured through Q-extraction procedures [2, 4, 6, 7]. In order to perform such measurements, it requires a glassy carbon and a platinum electrode in addition to an Ag/AgCl2 reference electrode. The redox state of exogenously added (1 μM) Q-1 or Q-2 is measured voltametrically (using a glassy carbon working electrode and a platinum electrode connected to an Ag/AgCl2 reference electrode). The working electrode is poised at
360 mV with respect to the reference electrode with a BAS CV-1B cyclic voltammeter. The outputs of the electrodes can be connected via an amperemeter to a standard recorder or computer for visualization and further analysis [2, 4]. Here, we describe in detail the isolation procedure of mouse heart mitochondria preparation for a simultaneous measurement of respiratory rates and Q pool redox states using customized electrodes.

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Materials Prepare all solutions using ultrapure water with a resistivity of 18.2 MΩ cm at 25  C and use reagents of analytical quality. Store reagents according to the manufacturer’s recommendations and carefully follow waste disposal regulations where applicable.

2.1 Isolation of Mouse Heart Mitochondria

1. Mitochondrial isolation buffer: 225 mM D-mannitol, 20 mM 3-(N-morpholino)propane sulfonic acid (MOPS), 75 mM sucrose, 1 mM ethylene glycol-bis(2-aminoethylether)-N,N, N0 ,N0 -tetraacetic acid (EGTA), 0.5 mM DL-dithiothreitol (DTT), pH 7.4. 2. Tissue homogenate digestion buffer: mitochondrial isolation buffer plus 0.05% nagarse (proteinase, bacterial, type XXIV). 3. Mitochondrial storage buffer: 225 mM D-mannitol, 20 mM MOPS, 75 mM sucrose, 0.1 mM EGTA, 75 mM potassium chloride (KCl), pH 7.4.

2.2 Assessing Respiratory Rates and Q Redox State

1. Respiration buffer: 120 mM D-mannitol, 20 mM MOPS, 5 mM potassium dihydrogen phosphate (KH2PO4), 60 mM KCl, 5 mM magnesium chloride (MgCl2), pH 7.4. 2. Tris–HCl 500 mM pH 7.4 (substrate dissolving buffer). Weight in 60.55 g Tris–HCl and add H2O ad 1000 mL. Carefully adjust pH by using 1 M NaOH (see Note 1). 3. Q-1 (e.g., 2,3-dimethoxy-5-methyl-6(3-methyl-2-butenyl)1,4-benzochinon, MW 250.29) or Q-2 (e.g., 2,3-dimethoxy5-methyl-6-geranyl-1,4-benzoquinone, MW 318.41), final concentration 1–2 μM. 4. Pyruvate (e.g., sodium pyruvate, MW 110.04). Use 55 mg pyruvate in 500 μL 500 mM Tris–HCl buffer to generate a 1 M stock solution (see Note 2). 5. Glutamate (e.g., L-glutamic acid monosodium salt hydrate, MW 169.11). Use 84.6 mg in 500 mM Tris–HCl buffer to generate a 1 M stock solution. Aliquot and store at
20  C. 6. Malate (e.g., L-(
)-malic acid, MW 134.09). Use 67.05 mg in 500 mM Tris–HCl buffer to generate a 1 M stock solution (see Note 1). Aliquot and store at
20  C. 7. Succinate (e.g., succinic acid disodium salt, MW 270.14). Use 135.05 mg in 500 mM Tris–HCl buffer to generate a 1 M stock solution (see Note 1). Aliquot and store at
20  C. 8. ADP (e.g., adenosine-50 -diphosphate disodium salt, MW 471.2). Use 4712 mg ad 20 mL 500 mM Tris–HCl buffer to generate a 500 mM stock solution and readjust the pH. Aliquot and store at
20  C.

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9. Rotenone (MW 394.42). Use 3 mg in 10 mL 96% ethanol to generate a 750 μM stock solution. Aliquot and store at
20  C. 10. Antimycin A (MW 532) (see Note 3). Use 3.5 mg in 1 mL 96% ethanol to generate a 6.5 mM stock solution. Aliquot and store at
20  C. 11. CAT (e.g., carboxyatractyloside potassium salt, MG770.82). Use 5 mg CAT ad 1.11 mL DMSO to generate a 5 mM stock solution. Aliquot and store at
20  C. 12. Azide (e.g., sodium azide, MW 65.01). Use 2.438 g in 10 mL H2O to generate a 3.75 M stock solution. Aliquot and store at
20  C.

3

Methods Carry out all procedures at temperature as specified.

3.1 Isolation of Mouse Heart Mitochondria

1. For the isolation of heart mitochondria, use 10–14-week-old mice. Remove hearts rapidly after decapitation and transfer to ice-cold mitochondrial isolation buffer. Carefully remove atria, right ventricle, connective tissue, and blood (see Note 4). Perform all further steps on ice (see Note 5). 2. Mince tissue with a scissor on a concave watch glass on ice to small pieces of approximately 1 mm3 and homogenize manually in ice-cold tissue homogenate digestion buffer using a glass-on-Teflon homogenizer until homogenous (see Note 6). No additional incubation time with nagarse is necessary (see Note 7). 3. Stop the action of nagarse by diluting the homogenate in 30 mL ice-cold mitochondrial isolation buffer. You may flip the solution gently for better mixing. Immediately move to step 4. 4. Centrifuge homogenate at 2000  g for 4 min at 4  C. 5. Pass supernatant through cheesecloth to separate homogenate from remaining particles (see Note 8). Immediately move to step 6. 6. Centrifuge flow-through at 12,000  g for 10 min at 4  C. 7. Resuspend the pellet in ice-cold mitochondrial storage buffer (see Note 9). 8. Determine mitochondrial protein content using, for instance, bicinchoninic acid assay with bovine serum albumin as a standard (see Note 10). 9. Always keep mitochondria on ice (see Note 11).

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Testing the Setup

295

1. Q-1 or Q-2 should become fully oxidized upon addition to the chamber in the presence of isolated mitochondria (in the absence of substrate but in the presence of 1 μM rotenone to prevent complex I substrates being oxidized) and fully reduced upon anaerobiosis. 2. 1 μM Q-1 or Q-2 should have no detectable effect upon either the state 4 respiration or the protonmotive force.

3.3 Assessing Respiratory Capacity and Q Redox State

1. Measure mitochondrial oxygen consumption and Q pool redox state at 30  C in air-saturated respiration buffer (200 nmol O2/mL at 95 kPa). 2. Add 0.03 mg heart mitochondria per mL (see Note 12) and 1 μL/mL of 1 mM Q-2. 3. Close the reaction vessel and wait until oxygen and Q traces have reached a steady-state phase (see Notes 13 and 14). 4. Add respiratory complex I substrates, i.e., 10 mM pyruvate, 10 mM glutamate, 2 mM malate (see Note 15). 5. Wait until oxygen and Q traces have reached a steady-state phase. 6. Add respiratory complex II substrate, i.e., 10 mM succinate (see Note 15). 7. Wait until oxygen and Q traces have reached a steady-state phase. 8. Add ADP to a final concentration of 2 mM (see Note 16). 9. Wait until oxygen and Q traces have reached a steady-state phase. 10. Add 1.5 μM of respiratory complex I inhibitor rotenone (see Note 17). 11. Wait until oxygen and Q traces have reached a plateau phase. 12. Add 2.5 μM of respiratory complex III inhibitor antimycin A (see Note 18). 13. Opt. 1, instead of using antimycin A, add 1–5 μM Cat, a highly selective and potent inhibitor of the adenine nucleotide translocator (ANT) (see Note 18). 14. Opt. 2, instead of using antimycin A, add 100 mM azide, an inhibitor of respiratory complex IV (COX) (see Note 18).

4

Notes 1. If dissolving an acidic substrate, add half the volume of Tris– HCl buffer 500 mM first and neutralize to pH 7.4 using 1 M NaOH (test with pH strip) before adding final volume.

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2. Pyruvate should always be diluted freshly and kept on ice before starting the experiment. It is possible, however, to weight in the powder and store it in appropriate tubes at 4  C. 3. Usually, antimycin A products represent an antimycin mixture with each product at different stoichiometries and a slightly different MW, e.g., antimycin A1 548.6; antimycin A2: 534.6; antimycin A3: 520.6; antimycin A4: 506.6. The exact amounts of each antimycin should be available from the respective Certificate of Analysis. Many researchers prefer to work with an approximate MW, here, for instance, 532. 4. Mitochondrial functions may differ depending on the developed blood pressure. Previously, we analyzed the high blood pressure heart muscle tissue of the left ventricle and septum and thus removed atria and right ventricle. 5. All tissues contain proteases and may degrade proteins and cell structures. If the conditions are given, all isolation steps should be carried out in the cold room at 4  C. 6. Naturally, the amount of buffer depends on the amount of tissue. As a rule of a thumb, 1 mL buffer suffices for the left ventricle and septum of a mouse heart. Furthermore, we recommend the use of a glass-on-Teflon potter with a volume of 2 mL. If a drilling machine is used for pottering, we recommend 800 rounds per minute (rpm). Note, this step may produce a lot of heat. Depending on the accuracy of tissue mincing, eight strokes are sufficient for homogenization. 7. The heart contains subsarcolemmal and interfibrillar mitochondria [8, 9]. The latter are pelleted with the myofibrils if the protease treatment is omitted. Of note, nagarse is suspected to damage mitochondria if applied for too long and thus other protocols have been established using trypsin and other proteases instead of nagarse [10]. In our hands and following this protocol, nagarse treatment resulted in perfectly coupled mitochondria. 8. Optionally, you may take dressing material (e.g., pure ‘Mull’ or nylon gaze) as used in clinical routine. 9. The mitochondrial pellet obtained by differential centrifugation often consists of two layers: a lower, more densely packed dark brownish layer, and an overlaying, poorly sedimented, pinkish layer of “fluffy” appearance [11]. The latter should be separated from the more densely packed mitochondria by gently snipping the centrifugation vessel. Optionally, you may carefully rinse the fluffy material away, e.g., by using a 1-mL pipette.

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10. You may use other methods to determine the protein content. Please note that some reagents may interfere with your readout, thereby falsifying the actual values. 11. Even if maintained on ice, isolated mitochondria will exhibit a decrease in respiratory activity over time (good quality for up to 2 h). Furthermore, mitochondria are stored on ice at relatively high concentration (approximately 1 mg/mL) and thus may become anoxic. To avoid undesirable side effects, one may flip the tube occasionally. 12. In our experience, 0.9 mg/mL is optimal for measuring mouse liver mitochondria. 13. To calibrate the Q electrode, fully oxidized Q (0% reduction, Qox) is taken as the steady-state achieved in the presence of 1 μM Q-1 or Q-2, mitochondria and 1 μM rotenone to inhibit endogenous NAD-linked dehydrogenases [2, 4, 6, 7]. With some mitochondria (such as purified plant mitochondria or some bacterial membranes [2, 6]), fully oxidized Q is apparent without addition of rotenone. 100% reduction (Qred) is achieved following anaerobiosis in the presence of a substrate such as succinate [12]. Although complete reduction can also be achieved in the presence of 1 mM KCN, anaerobiosis is the preferred method since it overcomes anomalies such as redox clamping such as with antimycin A. 14. The steady-state redox poise of the Q-pool reflects the equilibrium achieved between the dehydrogenases and the oxidases [7, 12–14]. Hence, the steady-state redox poise is substrate dependent. In other words, slow substrate oxidation rates (such as with malate/glutamate in the absence of ADP) results in a steady-state of 20–40% reduction, whereas rapid oxidation states (with succinate under non-phosphorylating conditions for instance) achieve steady-states in excess of 90% reduction levels. The engagement of the ATP synthase through the addition of ADP stimulates respiration which is reflected by an oxidation of the Q-pool often by as much as 20–30%. The addition of an uncoupler, however, collapses the mitochondrial membrane potential, stimulates respiration, and hence oxidizes the Q-pool to 10% Qred levels. For examples of steady-sytate redox levels, readers are referred to the following refs. 7, 12– 15. 15. The presence of respiratory substrates (and oxygen) in the absence of ADP generates a so-called LEAK respiration aka state 4. This is due to an attempt of mitochondria to rebuild the mitochondrial membrane potential against a proton leak across the inner membrane. This LEAK respiration is dependent on the substrate (combination) and a marker for the quality of the preparation.

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16. The presence of respiratory substrates (and oxygen) in the presence of ADP generates phosphorylating respiration aka state 3. Importantly, State 3 respiration converts to state 4 upon complete turnover of ADP to ATP. A prerequisite for this is that the amount of non-mitochondrial ATPase activity is low and mitochondria are not broken (another marker for the quality of the preparation) and that the added ADP can be turned over in a reasonable amount of time. Under the conditions described with a final concentration of 2 mM ADP, a sustained state 3 respiration is induced. If the conversion of states is the subject of investigation, a final concentration of 100 μM ADP or less is advisable. 17. Rotenone is highly lipophilic and thus easily enters cellular and mitochondrial membranes but also “contaminates” the wall of the reaction vessel. To avoid unintentional inhibition of respiratory complex I in follow-up experiments, thorough and repeated washing with 70% ethanol is required. Some researcher “wash” the reaction vessel with cell lysates or isolated liver mitochondria. 18. Generally, it is advisable to test respiratory inhibitors in a titration assay to define optimal concentrations and to avoid unintended side effects such as uncoupling. Furthermore, some inhibitors (such as salicylhydroxamic acid or azide) and substrates (ascorbate/TMPD, NADH) may interfere directly with the Q-electrode system, and hence, it is advisable to test in the absence of mitochondria.

Acknowledgments The authors thank Howard T. Jacobs for valuable discussions and financial support by the European Research Council (Advanced Grant 232738), the Academy of Finland (Centre of Excellence grant 272376 and Academy Professorship grant 256615), and the Tampere University Medical Research Fund (to H.T.J.). A.L.M. gratefully acknowledges funding support from University of Sussex and the BBSRC (BB/L022915/1 and BB/NO10051/ 1). C.V. acknowledges the kind support from Associazione Luigi Comini ONLUS and Telethon Foundation, Italy (grants GGP19007, 23706). References 1. Moore AL, Bonner WD, Rich PR (1978) The determination of the proton-motive force during cyanide-insensitive respiration in plant mitochondria. Arch Biochem Biophys 186: 298–306

2. Dry IB, Moore AL, Day DA, Wiskich JT (1989) Regulation of alternative pathway activity in plant mitochondria: nonlinear relationship between electron flux and the redox poise

Mitochondrial Q Pool Redox State of the quinone pool. Arch Biochem Biophys 273:148–157 3. Perales-Clemente E, Bayona-Bafaluy MP, Pe´rez-Martos A et al (2008) Restoration of electron transport without proton pumping in mammalian mitochondria. Proc Natl Acad Sci U S A 105:18735–18739. https://doi.org/ 10.1073/pnas.0810518105 4. Moore AL, Dry IB, Wiskich JT (1988) Measurement of the redox state of the ubiquinone pool in plant mitochondria. FEBS Lett 235: 76–80. https://doi.org/10.1016/0014-5793 (88)81237-7 5. Lemieux H, Blier PU, Gnaiger E (2017) Remodeling pathway control of mitochondrial respiratory capacity by temperature in mouse heart: electron flow through the Q-junction in permeabilized fibers. Sci Rep 7:2840. https:// doi.org/10.1038/s41598-017-02789-8 6. Zannoni D, Moore AL (1990) Measurement of the redox state of the ubiquinone pool in Rhodobacter capsulatus membrane fragments. FEBS Lett 271:123–127. https://doi.org/10. 1016/0014-5793(90)80387-x 7. Van den Bergen CW, Wagner AM, Krab K, Moore AL (1994) The relationship between electron flux and the redox poise of the quinone pool in plant mitochondria. Interplay between quinol-oxidizing and quinonereducing pathways. Eur J Biochem 226: 1071–1078. https://doi.org/10.1111/j. 1432-1033.1994.01071.x 8. Palmer JW, Tandler B, Hoppel CL (1977) Biochemical properties of subsarcolemmal and interfibrillar mitochondria isolated from rat cardiac muscle. J Biol Chem 252:8731–8739

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9. Weinstein ES, Benson DW, Fry DE (1986) Subpopulations of human heart mitochondria. J Surg Res 40:495–498 10. Fannin SW, Lesnefsky EJ, Slabe TJ et al (1999) Aging selectively decreases oxidative capacity in rat heart interfibrillar mitochondria. Arch Biochem Biophys 372:399–407. https://doi.org/ 10.1006/abbi.1999.1508 11. Laird AK, Nygaard O, Ris H, Barton AD (1953) Separation of mitochondria into two morphologically and biochemically distinct types. Exp Cell Res 5:147–160. https://doi. org/10.1016/0014-4827(53)90100-1 12. Affourtit C, Krab K, Leach GR et al (2001) New insights into the regulation of plant succinate dehydrogenase. On the role of the protonmotive force. J Biol Chem 276: 32567–32574. https://doi.org/10.1074/jbc. M103111200 13. Affourtit C, Albury MS, Krab K, Moore AL (1999) Functional expression of the plant alternative oxidase affects growth of the yeast Schizosaccharomyces pombe. J Biol Chem 274: 6212–6218. https://doi.org/10.1074/jbc. 274.10.6212 14. Affourtit C, Krab K, Moore AL (2001) Control of plant mitochondrial respiration. Biochim Biophys Acta 1504:58–69 15. Wagner AM, Krab K, Wagner MJ, Moore AL (2008) Regulation of thermogenesis in flowering Araceae: the role of the alternative oxidase. Biochim Biophys Acta 1777:993–1000. https://doi.org/10.1016/j.bbabio.2008. 04.001

Chapter 20 Experimental Setup for Investigation of Acute Mitochondrial Oxygen Sensing in Primary Cells Fenja Knoepp, Norbert Weissmann, Natascha Sommer, and Marten Szibor Abstract The ability to sense and respond to acute changes in oxygen is essential for the viability of cells and organisms. To study molecular mechanisms of acute oxygen sensing, we established a setup for the adjustment of acute hypoxic conditions in cultured cells, exemplified here for the use of primary pulmonary arterial smooth muscle cells (PASMCs). The mitochondrial electron transport chain (ETC) is the main consumer of oxygen but recently also emerged as essential oxygen sensor suggesting that the ETC itself adapts its electron flux to oxygen availability. To test this assumption and to experimentally manipulate electron flux through the ETC, we used alternative oxidase (AOX), which bypasses the cytochrome pathway of the ETC when blocked. The described combination of our experimental setup and AOX allowed us in previous publications unprecedented insights into the role of the ETC in cellular oxygen sensing and cellular response mechanisms in living cells. Against this background, we here describe and discuss this method in detail, which will allow transfer to other cell types and research questions. Key words Lung, Mitochondria, Acute hypoxia, Oxygen sensing, Alternative oxidase (AOX), Smooth muscle cells

1

Introduction The ability to sense and respond to acute changes of oxygen concentration is essential for the viability of cells and organisms. Different molecular mechanisms ruling adaptational processes to oxygen availability are currently debated [1]. We and others suggested that mitochondria and specifically the mitochondrial electron transport chain (ETC) play an essential role in this process [2– 5]. Unfortunately, adaptation to oxygen availability requires an integrated cell response, making in vitro assays less informative. To study molecular mechanisms of acute oxygen sensing on the cellular level, we established an experimental setup for the adjustment of acute hypoxic conditions, exemplified here for the use of pulmonary arterial smooth muscle cells (PASMC). We here describe in detail this setup that was previously successfully applied

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_20, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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by our group [2, 5]. As a tool to manipulate electron flux through ETC, we used alternative oxidase (AOX) (Fig. 1) [7, 8]. The ETC is the main consumer of oxygen but recently emerged also as a central oxygen sensor in isolated PASMC [2–5]. We, for instance, demonstrated that signals arising from the ETC itself adapt electron flux to oxygen availability, which in effect then coordinates cell responses [2, 5]. Mechanistically, we found that a reduced ETC triggers the production of reactive oxygen species (ROS) prior to the increase in cellular calcium and contraction [5]. The observed mitochondrial ROS signaling is likely at the very tip in the hierarchy of a signaling cascade underlying the redirection of pulmonary capillary blood flow toward alveolar areas of high oxygen partial pressure (hypoxic pulmonary vasoconstriction, HPV), a physiologic response process also known as the von Euler-Liljestrand mechanism. An essential aspect toward this conclusion [2, 5] was to establish and validate an experimental setup for acute oxygen adjustments in living cells and the use of AOX, an ubiquinol oxidase which introduces a branch point into the respiratory electron transport chain, thereby bypassing the cytochrome pathway of the ETC when blocked (Fig. 1) [9]. AOX is present in plants, various fungi, and microorganisms, but not in insects and mammals, and provides a bypass of ETC complexes III and IV when impaired [9]. Concomitant with branching the ETC, AOX prevents excessive superoxide production and thus interferes with ETC signaling [5, 6, 9, 10]. To use AOX, we took advantage of a recently generated mouse strain (AOXRosa26) that ubiquitously expresses AOX originating from Ciona intestinalis [6]. The combination of acute oxygen adjustments and AOX allowed us to study the interplay between oxygen availability and ETC-related signaling and its impact on the integrated cellular response with unprecedented depth. The described system for induction of precisely controlled oxygen concentrations was used successfully by us for measurement of mitochondrial redox state by Raman spectroscopy or cellular membrane potential by patch clamp in PASMC previously [2, 5]. However, the setup is broadly applicable for many cell types and microscopy methods. Taken together, we describe here all the crucial steps from sample preparation to the construction of a customized perfusion system for the application of acute hypoxia to living cells. We also discuss potential pitfalls in studying the role of the ETC and its constituents in oxygen-related signal transduction and provide an overview on AOX expression as tool to gain insight into the role of the ETC in oxygen sensing.

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Fig. 1 Expression of AOX serves as tool to investigate oxygen sensing [5]. Inhibition of the ETC at complex IV (e.g., by hypoxia) reduces the redox state of upstream constituents of the ETC. As a consequence, the NADH/ NAD+ ratio may increase, succinate may accumulate, and excessive amounts of reactive oxygen species (ROS) may be produced. Alternative oxidase (AOX) is a single protein, non-protonmotive respiratory enzyme that acts as a sink for electrons from a reduced quinol pool and directly decreases oxygen to water when the cytochrome pathway of the ETC is blocked [6]. Thereby, AOX abolishes hypoxia-induced accumulation of electrons, oxidizes substrates of the ETC, and decreases excessive ROS production. AOX has proven effective in manipulating hypoxia-induced signaling and hypoxia-related cellular downstream responses

2

Materials Important note: prepare all solutions using ultrapure water and reagents of analytical quality. Prepare and store all reagents as specified by the manufacturer. When disposing of waste materials, carefully follow all waste disposal regulations.

2.1 Laboratory Equipment

• Inverse microscope (e.g., DMI3000b, Leica Microsystems, Wetzlar, Germany), equipped with a microscope camera (e.g., ProgRes® FM, Jenoptik, Jena, Germany), a 40-fold magnification objective (e.g., LUCPLFLN 40, Olympus, Tokyo, Japan), and, if applicable, the possibility to perform fluorescence measurements. • Personal computer with operation software for the microscope camera (e.g. ProgRes® CapturePro Software, Jenoptik, Jena, Germany). • Microscale. • Magnetic stirrer. • pH meter.

2.2

Cells

• Pulmonary arterial smooth muscle cells that are grown on CELLview™ cell culture dishes with glass bottom (35 mm,

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627860, Greiner Bio-One, Frickenhausen, Germany) and cultured in Smooth muscle cell growth medium 2 plus supplements (SMC-medium 2, C-22062, PromoCell, Heidelberg, Germany). 2.3

Chemicals

• Sodium chloride St. Louis, USA).

(NaCl,

31434,

Sigma-Aldrich,

• Potassium chloride (KCl, 5346.1, Carl Roth, Karlsruhe, Germany). • Calcium chloride (CaCl2, CN93.1, Carl Roth, Karlsruhe, Germany). • Magnesium chloride (MgCl2, KK36.1, Carl Roth, Karlsruhe, Germany). • Sodium dihydrogen phosphate dihydrate (NaH2PO4
2H2O, 1063421000, Merck Millipore, Darmstadt, Germany). • Sodium hydrogen carbonate (NaHCO3, 8551, Carl Roth, Karlsruhe, Germany). •

D(+)-Glucose

(X997.1, Carl Roth, Karlsruhe, Germany).

• Normoxic gas mixture (21% O2, 5% CO2, rest N2, 592091, custom-made by Praxair, Danbury, CT, USA). • Hypoxic gas mixture (1% O2, 5% CO2, rest N2, 650329, custom-made by Praxair, Danbury, CT, USA). 2.4 Perfusion and Temperature Control

• Valve-controlled gravity-driven perfusion system (e.g., VC38xG Series, ALA Scientific Instruments, Farmingdale, New York, USA). • Bubbler manifold for Perfusion System (e.g., ALA-Bubbler 8, ALA Scientific Instruments, Farmingdale, New York, USA). • Glass syringes as perfusion reservoirs (50 ml, Luer Lock tip, e.g., C692.1, Carl Roth, Karlsruhe, Germany). • Dual channel heater controller (e.g., TC-344C, Warner Instruments, Hamden, CT, USA). • Multi-line in-line solution heater (e.g. SHM-8, Warner Instruments, Hamden, CT, USA). • Cell culture dish microincubator (DH-40iL microincubator, Warner Instruments, Hamden, CT, USA). • Perfusion insert (RC-37F, Warner Instruments, Hamden, CT, USA). • Vacuum grease (e.g., Dow Corning® 111 Valve Lubricant & Sealant, 64-0275, Warner Instruments, Hamden, CT, USA). • Vacuum Waste Kit (VWM, ALA Scientific Instruments, Farmingdale, New York, USA).

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• Precision restrictor fitting (e.g., R-008-6, AirCom Pneumatic GmbH, Ratingen, Germany). • Silicone and airtight Tygon® tubes of different diameters. • Three-way stopcocks (e.g. Discofix™ 3-way Stopcock, B. Braun, Melsungen, Germany). • Tubing and Connector Kit (e.g. 64-1565, Warner Instruments, Hamden, CT, USA). 2.5

Consumables

• Spatula and weighing dishes. • Beaker (filling volume ~ 1200 ml). • Magnetic stirrer rod. • Bubbler stones (e.g., S KU: 01-40, AutoMate Scientific, Berkeley, California USA). • Calibrated volumetric flask (filling volume 1000 ml). • Laboratory glass bottle (filling volume 1000 ml). • Squeeze bottle. • Ultrapure water. • Coverslips (~5 cm in diameter). • Pipette (1 ml) with tips. • Cotton buds. • Tissues.

3

Methods Carry out all procedures at temperatures as specified. 1. Use clean and autoclaved glassware for the preparation of Tyrode’s solution.

3.1

Preparations

1. Prepare 1 l of Tyrode’s solution: (a) Weigh in 7.4 g NaCl, 0.4 g KCl, 0.06 g NaH2PO4, 0.21 g MgCl2, 0.2 g CaCl2, 1.85 g NaHCO3 and 1.8 g glucose (see Note 1). (b) Transfer the weighed chemicals into a beaker. (c) Add a clean magnetic stirrer rod of appropriate size. (d) Add ~900 ml of ultrapure water (see Note 2). (e) Place the beaker on top of a magnetic stirrer and switch the stirrer on (see Note 3). (f) Using a bubbler stone, aerate the solution inside the beaker with a gas mixture of 21% O2, ~5% CO2, rest N2 for at least 10 min (see Note 4).

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(g) Remove and thoroughly clean the bubbler stone (see Note 5). (h) Pour the Tyrode’s solution into a calibrated volumetric flask (1000 ml) (see Note 6). (i) Wash off the remaining Tyrode’s solution in the beaker with ultrapure water (e.g., via a squeeze wash bottle) and add this solution to the Tyrode’s solution in the calibrated volumetric flask. (j) Fill the volumetric flask exactly up to the mark with ultrapure water (see Note 7). (k) Transfer the Tyrode’s solution into a fresh glass bottle and keep at room temperature until use (see Note 8). 2. Install a valve-controlled gravity-driven perfusion system whose reservoirs (i.e., glass syringes) can be separately gassed via the bubbler manifold (see Note 9). 3. Connect the reservoirs of the perfusion system with a multi in-line solution heater (see Note 10) via separate airtight Tygon tubes (see Note 11). 4. Place the cell culture dish microincubator onto the microscope stage and securely fix it via the supplied screws (see Note 12). 5. Plug the in-line heater and the culture dish microincubator in the dual channel heater controller and pre-warm both devices to 37  C. 6. Fill two reservoirs with the Tyrode’s solution (prepared in step 1). 7. Aerate one of the reservoirs with normoxic gas mixture (21% O2, 5% CO2, rest N2) and the second one with hypoxic gas mixture (1% O2, 5% CO2, rest N2). 8. Open one valve and test the fluid velocity, which should not exceed 1 ml/min (see Note 13). 9. Close all valves. 10. Check the fluid level inside the reservoirs and top off with Tyrode’s solution, if necessary (see Note 14). 11. Connect the suction device (supplied with the perfusion insert) with the vacuum pump via suitable silicone tubes. 3.2 Experimental Procedure

1. Prepare the perfusion insert (Fig. 2a) by applying a very thin layer of vacuum grease to the bottom edge. It is extremely important to verify that the access path for inflow is not blocked by the vacuum grease (see Note 15). 2. Carefully press a coverslip onto the silicone (see Note 16). 3. Again, verify that the inflow is not blocked by vacuum grease. 4. Set the perfusion insert with attached coverslip aside until use.

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Fig. 2 Perfusion assembly for hypoxic measurement conditions in cell culture dishes [5]. (a) (1) CELLview™ cell culture dishes with glass bottom (35 mm, 627860, Greiner Bio-One) and (2) the Open perfusion dish insert (RC-37W Perfusion, Warner Instruments) that is assembled with fluid (3) inlet and (4) outlet ports and connected to an (5) in-line solution heater. (b) Assembled Measurement chamber. To control the velocity of fluid application, the Tyrode’s Solution is delivered to the perfusion inlet port through a (6) precision restrictor fitting (R-008-6, AirCom Pneumatic GmbH, Ratingen, Germany) before being removed from the chamber by means of (7) a height adjustable aspirator. (c) For keeping the cells at a physiological temperature, the assembled measurement chamber is placed in (8) a cell culture dish microincubator (DH-40iL microincubator, Warner Instruments, Hamden, CT, USA)

5. Take one cell culture dish containing previously isolated PASMCs from the cell culture incubator (see Note 17). 6. Aspirate off the cell culture medium via a 1-ml pipette (see Note 18).

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7. To avoid draining of the PASMC, the following steps (see steps 8–16) must be performed quickly and without delay. 8. Use a cotton bud to thoroughly dry the periphery surrounding the glass bottom of the culture dish but be careful to not touch the glass bottom (see Note 19). 9. Carefully remove the coverslip from the perfusion insert (see step 4). 10. Place the perfusion insert into the cell culture dish (Fig. 2b). 11. Install the assembled culture dish with perfusion insert in the culture dish incubator (Fig. 2c) and properly fix it via the supplied clamps (see Note 20). 12. Connect the perfusion insert’s inflow with the in-line solution heater via a Tygon tube. 13. Insert the assembled suction device into the specified place of the perfusion insert. 14. Start the vacuum pump for aspiration. 15. Open the valve for Normoxic Tyrode’s solution. 16. Ensure consistent flow of the Tyrode’s solution inside the perfusion insert. 17. Use the 40 magnification to focus on the PASMC of interest. 18. Start the normoxic experiment (e.g., Raman spectroscopy, patch clamp, fluorescence measurements, calcium imaging or other). 19. Switch the perfusion to hypoxic Tyrode’s solution. 20. Wait for 2–3 min to establish stable hypoxic conditions inside the section of interest (see Note 21). 21. Start hypoxic experiment (e.g., Raman spectroscopy, patch clamp, fluorescence measurements, calcium imaging or other). 22. Stop the perfusion upon acquisition. 23. Remove the culture dish, thoroughly clean the perfusion insert, and prepare it for the next experiment (see steps 1–4). 24. For normoxic control experiments, repeat steps 5–23 with fresh PASMC but do not switch to hypoxic conditions (as described under step 19). Instead, use normoxic Tyrode’s solution. 25. For statistical analysis, repeat normoxic and hypoxic experiments with at least four culture dishes with PASMC from the different study groups. Here, we used AOX-transgenic and wild-type littermates to manipulate the ETC and test its effect on acute oxygen sensing (Fig. 1).

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Notes 1. Depending on the supplier of chemicals, the molecular weight may vary, which requires an adjustment of the amounts. The final concentration should be 126.7 mM NaCl, 5.4 mM KCl, 0.42 mM NaH2PO4, 1.05 mM MgCl2, 1.8 mM CaCl2, 22 mM NaHCO3, and 10 mM glucose. 2. As the ionic composition, pH and osmolarity needs to be carefully controlled, it is crucial to use ultrapure water, which is essentially free of ions, organic compounds, heavy metals, endotoxins, and bacteria. 3. The salts will dissolve very quickly, resulting in a clear liquid. 4. This procedure will adjust the pH to ~7.4. You can use a pH-meter to verify the physiological pH-value. 5. To avoid contaminations, e.g., as remaining glucose may induce mold growth, properly clean the bubbler stone by aeration of ultrapure water, followed by thorough rinsing and subsequent air drying. 6. To avoid spillage, it is recommendable to use a funnel. 7. To prepare the Tyrode’s solution with specified ionic composition, the total volume should be exactly 1000 ml. 8. You may keep the solution at 4  C but for no more than 1 week. 9. Make sure that the syringes, which serve as reservoirs, are equipped with three-way stopcocks. This allows to prevent leakage, e.g., during preparations or washing procedures at the end of experiments. 10. In-line heaters allow warming of the applied fluid directly before application to the measurement chamber. Make sure to thoroughly occlude all unused channels of the in-line heater. 11. Choose tubes that are flexible enough to be squeezed by the perfusion system’s valves. 12. Dependent on the microscope, the microincubator may require the additional purchase of a special stage adapter. 13. If fluid velocity is too high, reduce the height of the perfusion system’s reservoirs and/or include a precision restrictor fitting into the tube connecting the in-line heater with the perfusion chamber. 14. It is not recommendable to top off the Tyrode’s solution inside the reservoirs during experiments. 15. Silicone is required to later form a seal between the perfusion insert and the cell culture dish, thereby preventing fluid leakage and enabling a laminar flow inside the measurement section. Therefore, it is crucial that the whole bottom of the perfusion insert is surrounded with silicone.

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16. This will ensure an even distribution of the vacuum grease on the bottom of the perfusion insert. 17. From this point onward, sterile cell culture conditions are not required any more. All steps can be performed at a non-sterile bench next to microscopy, e.g., Raman spectroscopy. 18. The PASMC growth medium contains phenol red whose fluorescence signal would hinder certain applications such as Raman signals. Therefore, if applicable, the medium needs to be replaced by Tyrode’s solution during experiments. 19. This step will promote sealing between the perfusion insert and the culture dish via the previously applied silicone grease. 20. As maintaining a consistent optical focus throughout the experiment is essential for microscopy techniques or to obtain good-quality Raman spectra, the cell culture dish needs to be sufficiently secured. 21. To specify the precise oxygen concentration in close proximity to the PASMC, an optical oxygen sensor can additionally be placed into the measurement section.

Acknowledgments We thank Dr. Monika Brosien for critical reading of the manuscript and Susanne Lich for technical advice and support. This work was supported by Deutsche Forschungsgemeinschaft (DFG, German Research Foundation)—Project number 268555672—SFB 1213, Project A06 and the Excellence Cluster Cardio-Pulmonary System (CPI). References 1. Sommer N, Strielkov I, Pak O, Weissmann N (2016) Oxygen sensing and signal transduction in hypoxic pulmonary vasoconstriction. Eur Respir J 47:288–303. https://doi.org/ 10.1183/13993003.00945-2015 2. Sommer N, Hu¨ttemann M, Pak O et al (2017) Mitochondrial complex IV subunit 4 isoform 2 is essential for acute pulmonary oxygen sensing. Circ Res 121:424–438. https://doi.org/ 10.1161/CIRCRESAHA.116.310482 3. Dunham-Snary KJ, Wu D, Sykes EA et al (2017) Hypoxic pulmonary vasoconstriction: from molecular mechanisms to medicine. Chest 151:181–192. https://doi.org/10. 1016/j.chest.2016.09.001 4. Smith KA, Schumacker PT (2019) Sensors and signals: the role of reactive oxygen species in hypoxic pulmonary vasoconstriction. J Physiol

597:1033–1043. https://doi.org/10.1113/ JP275852 5. Sommer N, Alebrahimdehkordi N, Pak O et al (2020) Bypassing mitochondrial complex III using alternative oxidase inhibits acute pulmonary oxygen sensing. Sci Adv 6:eaba0694. https://doi.org/10.1126/sciadv.aba0694 6. Szibor M, Dhandapani PK, Dufour E et al (2017) Broad AOX expression in a genetically tractable mouse model does not disturb normal physiology. Dis Model Mech 10:163–171. https://doi.org/10.1242/dmm.027839 7. Perales-Clemente E, Bayona-Bafaluy MP, Pe´rez-Martos A et al (2008) Restoration of electron transport without proton pumping in mammalian mitochondria. Proc Natl Acad Sci U S A 105:18735–18739. https://doi.org/ 10.1073/pnas.0810518105

Mitochondria in Acute Hypoxia 8. Rustin P, Jacobs HT (2009) Respiratory chain alternative enzymes as tools to better understand and counteract respiratory chain deficiencies in human cells and animals. Physiol Plant 137:362–370. https://doi.org/10. 1111/j.1399-3054.2009.01249.x 9. Szibor M, Gainutdinov T, Fernández-Vizarra E et al (2020) Bioenergetic consequences from xenotopic expression of a tunicate AOX in

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mouse mitochondria: switch from RET and ROS to FET. Biochim Biophys Acta Bioenerg 1861:148137. https://doi.org/10.1016/j. bbabio.2019.148137 10. Robb EL, Hall AR, Prime TA et al (2018) Control of mitochondrial superoxide production by reverse electron transport at complex I. J Biol Chem 293:9869–9879. https://doi. org/10.1074/jbc.RA118.003647

Chapter 21 Assessing the Redox Status of Mitochondria Through the NADH/FAD2+ Ratio in Intact Cells Haoyu Chi, Gauri Bhosale, and Michael R. Duchen Abstract This section aims to describe the measurement of NADH and FAD2+ levels in intact cells using fluorescence microscopy. Both NADH and FADH2 are major electron donors for the electron transport chain through shifting of their redox status. Furthermore, within their redox couples, only NADH and FAD2+ are fluorescent. Therefore, calibration of the NADH and FAD2+ fluorescence signal is a crucial factor in accurately assessing mitochondrial function and redox status. Key words NADH, FAD2þ, Autofluorescence, Redox status, ETC

1

Introduction Mitochondria are often referred to as the “powerhouse” of the cell, as they generate ATP through the function of the electron transport chain (ETC). The ETC is composed of several complexes which use energy derived from redox reactions to pump protons across the inner mitochondrial membrane to generate an electrochemical gradient. This proton motive force is subsequently harnessed to phosphorylate ADP to ATP by the F1F0-ATP synthase. Within the mitochondrial electron transport chain (ETC), Complex I (NADH: ubiquinone oxidoreductase) and Complex II (Succinate dehydrogenase) utilize NADH and FADH2, respectively, as substrates/coenzymes. Through oxidation, these molecules donate protons and electrons to the ETC, therefore contributing to the establishment of the mitochondrial membrane potential and ATP production [1]. Consequently, NADH (nicotinamide adenine dinucleotide) and FAD2+ (flavin adenine dinucleotide) molecules are shifting constantly between the oxidized and reduced forms, due to the function of the different enzymes implicated in energy-producing pathways, for example, matrix dehydrogenases in the citric acid

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cycle. The balance between the two distinct forms is crucial for determining redox status and ETC function. The reduced form of NAD+ (NADH) and the oxidized form of FAD (FAD2+) are both intrinsically fluorescent, with distinct spectral properties, while their redox pair is not fluorescent. Thus, we can measure both species simultaneously using a confocal microscope. This allows us to assess the redox status of the mitochondrial NADH and FAD2+ pools [2]. To determine the dynamic range of NADH and FAD2+ pools, drugs including NaCN and FCCP (phospho-trifluoromethoxy carbonyl cyanide phenylhydrazone) are often used. NaCN is an inhibitor of complex IV (COX: cytochrome c oxidase); therefore, it prevents the reduction of oxygen to water. This in turn inhibits the transfer of electrons through the ETC. This downstream inhibition will inhibit complex I and II, resulting in the accumulation of NADH and FADH2, thus providing a measure for maximally reduced mitochondrial NADH and FADH2 pools. Meanwhile, FCCP is an uncoupler of ETC, which allows protons to travel freely across the mitochondrial inner membrane independent of the ATP synthase, resulting in maximum activity of ETC complexes, which are normally constrained by the proton gradient. In this case, NAD+ and FAD2+ accumulate in their oxidized form due to rapid oxidation by CI and CII, and therefore gives a measure of the maximally oxidized pool. Taken together, these drug treatments allow us to determine the dynamic range of the NADH and FADH2 pools shifting between the extremes of their redox states [3, 4]. Overall, this approach is to investigate redox status of NADH and FADH2 pools within their maximum and minimum dynamic range, thus allowing us to extrapolate backwards to define a resting “redox index” as a measure which is defined by the balance of resting respiratory rate and resting generation of reduced cofactors by the TCA cycle. These measurements are taken using a confocal microscope and application of drugs (NaCN and FCCP).

2 2.1

Materials Equipment

1. Cells need to be seeded into a fluorescence imaging compatible coverslips, plates, or dishes; it is very important to ensure that the imaging glass is UV-light transmissible for the NADH signal. 2. A fluorescence microscope with excitation in the UV-range (normally containing a 355 nm laser). 3. It is necessary to use UV compatible objectives. Recording medium: Use a recording medium that is compatible with your cells, ideally the medium used to grow the cells without phenol red and additional pH buffers such as HEPES.

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Drugs: NaCN and FCCP are required to generate maximum and minimum pools of NADH or FAD2+. NaCN solutions should be prepared fresh on the day of the assay and should be pH adjusted. NaCN is volatile, and concentrations decrease rapidly after preparation, and so it must be used fresh. 1 mM FCCP stock solutions can be prepared and stored at 20  C.

3 3.1

Methods Imaging

1. Prepare the drugs: Prepare a 10 solution of NaCN in recording medium and a 10 FCCP stock in recording medium, vortex to dissolve, and keep on ice (see Note 1). 2. Set up the microscope. The two optical paths required are: (a) NADH: excitation 450 nm  30 nm.

at

355

nm,

emission:

(b) FAD2+ signal: excitation at 458 nm (if you do not have a line at 458, it is possible to use the 488 nm line of an Argon laser), emission: 550 nm  50 nm (see Note 2). 3. Aspirate the growth medium from the cells and wash once with pre-warmed recording medium before replacing with recording medium. Proceed to the microscope. 4. Set up the stage and focus. Find and save optimum image positions to image an appropriate number of cells (see Note 3). 5. Adjust microscope settings (see Note 4). 6. We recommend trying time series at first on the microscope to record the change in fluorescence signal throughout the experiment, also helps with identifying time required to equilibrate after drug additions (see Note 5). 7. Start the acquisition to take images of cells under basal conditions with no drugs (we recommend 3–5 timepoints) (see Note 6). 8. Add NaCN, according to previously determined concentrations. Allow to equilibrate and start acquisition (see Note 7). 9. Aspirate the recording medium after the acquisition has finished and carefully wash the dish/plate to remove the NaCN (see Note 8). 10. Add fresh recording medium. Add the FCCP into the Fluorodish or imaging plate, again wait for the drug to work and start the acquisition (see Note 9). 3.2

Data Analysis

1. Quantification: Use an imaging processing software such as ImageJ to quantify the NADH/FAD2+ signal from each cell under the different conditions (see Notes 10 and 11).

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2. Determine the status of NADH/FAD2+ within their respective dynamic range and calculate the pool index of NADH (reduced states)/FAD2+ (oxidized states) using this formula: Pool Index ¼

4

F basal  F min F max  F min

Notes 1. For most cells, the final concentration of NaCN required to maximally inhibit respiration is between 0.1 and 10 mM, and the final effective concentration of FCCP is 1–5 μM. Drug concentrations need to be optimized for individual cell lines. 2. If possible, set up a bright field path as well to confirm the outline of the cells. 3. If available, record the position of several fields on the microscope software to increase sampling from each condition; this would maximize the number of cells imaged. 4. Set the laser power or exposure as low as possible to limit photo-toxicity or bleaching, while ensuring you obtain a good signal: noise ratio. Adjust the master gain and digital offset accordingly if available to obtain an optimum signal output. These signals are dim—on most microscope systems we find we need to maintain a wide pinhole on a confocal microscope to maximize signal detection. We also recommend setting the incubation of either the microscope chamber or the stage at 37  C beforehand, to keep the temperature constant during the imaging process. 5. We recommend taking one time series at least once per cell line/type, to determine exactly how long it takes for the signal to reach plateau (maximum and minimum) after the drug addition. 6. Number of images depends on the confluency and homogeneity of cell population, generally at least 3–5 fields per dish/plate cells. 7. We would advise to try a range of drug concentrations, utilizing information from the time series to ensure you are capturing the maximum/minimum intensity post drug application. 8. A Pasteur Pipette with a thin tip is recommended for washing and aspirating the medium after the NaCN addition. It is very crucial to keep the dish/plate steady on the stage because we need to be able to find the previously saved positions. 9. For the drug additions, there are several things worth considering. Apart from the duration and dosage of drugs, the order of drug additions can also be adjusted with different cell types.

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Some cells are more sensitive to NACN and do not exhibit a robust NADH reduction post FCCP addition. Therefore, it is possible to first add FCCP, followed by NaCN without a wash step. 10. We recommend using Image J FIJI. Draw a region of interest around each cell, apply appropriate thresholding and record the signal intensity, subtract the background signal, obtain the basal (Fbasal), maximum (Fmax), and minimum signal (Fmin). Note that the intensity of NADH and FAD2+ should be inversely correlated with each drug treatments, for example, NADH pool reaches the maximum under NaCN addition, whereas FAD2+ reaches minimum here (Fig. 1).

Fig. 1 NAD(P)H autofluorescence in human fetal fibroblasts. (a) At baseline, NAD(P)H autofluorescence intensity is primarily observed in the mitochondrial network. (b) After the addition of NaCN, the mitochondrial NAD(P)H autofluorescence should increase as the NAD(P)H pool is maximally reduced due to the inhibition of Complex IV. (c) In the presence of the uncoupler FCCP, NAD(P)H is maximally oxidized due to increased activity of the ETC, resulting in a significant reduction in fluorescence intensity. (d) Quantification of NAD(P)H fluorescence units (F.U.), where baseline NAD(P)H pool is calculated relative to the maximally reduced and maximally oxidized fluorescence values. Scale bar: 20 μm The calculation of pool index of the two cells: F min Pool Index ¼ FFbasal ¼ 420279 573279 ¼ 0:48 max F min F min ¼ 406278 Pool Index ¼ FFbasal 523278 ¼ 0:50 max F min

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11. It is important to note that the autofluorescence signal will originate from both NADH and NADPH pools within the cell, which can only be spectrally distinguished using fluorescence lifetime imaging. Therefore, often NADH is referred to as NAD(P)H when quantifying autofluorescence.

Acknowledgments Funding for assay development and manuscript preparation was provided by the BBSRC and Mundipharma. References 1. Lehninger AL, Nelson DL, Cox MM (2013) Lehninger principles of biochemistry, 6th edn. W.H. Freeman, New York 2. Blacker TS, Duchen MR (2016) Investigating mitochondrial redox state using NADH and NADPH autofluorescence. Free Radic Biol Med 100:53–65

3. Duchen MR, Surin A, Jacobson J (2003) Imaging mitochondrial function in intact cells. Methods Enzymol 361:353–389 4. Bartolome F, Abramov AY (2015) Measurement of mitochondrial NADH and FAD autofluorescence in live cells. Methods Mol Biol 1264: 263–270

Chapter 22 Monitoring Mitochondrial Membrane Potential in Live Cells Using Time-Lapse Fluorescence Imaging Gabriel Esteban Valdebenito and Michael R. Duchen Abstract The mitochondrial membrane potential (ΔΨm) generated by proton pumps (Complexes I, III, and IV) is an essential component in the process of energy generation during oxidative phosphorylation. Tetramethylrhodamine, methyl ester, perchlorate (TMRM) is one of the most commonly used fluorescent reporters of ΔΨm. TMRM is routinely employed in a steady state for the measurement of membrane potential. However, it can also be utilized with time-lapse fluorescence imaging to effectively monitor the changes in membrane potential in response to a given stimulus by analyzing the change in distribution of the dye with time. Key words Mitochondria membrane potential, TMRM, Fluorescence microscopy, Uncoupler, Primary skin fibroblasts

1

Introduction The mitochondrial membrane potential (ΔΨm) is a semi-quantitative read-out for the full proton-motive force defined as the difference in electrical potential between the mitochondrial matrix and the cytosol [1]. Alteration in ΔΨm can indicate disruption in oxidative phosphorylation, ATP synthesis, or ionic exchange through the mitochondrial membrane [2, 3]. TMRM, tetramethylrhodamine, ethyl ester (TMRE), rhodamine 123 (Rh123), and JC1 are cationic, membrane permeant reporters that are commonly used to measure mitochondrial membrane potential [4–6]. These dyes are able to cross the cell membrane and partition between compartments depending on the electrochemical potential gradients [2]. The magnitude of the accumulation of these dyes in negatively charged compartments is described by the Nernst equation [2] but is also affected by the mitochondrial binding of the probe. TMRM is widely used due to

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_22, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Equilibration of 25 nM TMRM in the mitochondria after 30 min of incubation in skin fibroblasts. Nuclei were labeled with Hoechst 33342

Fig. 2 Distribution of TMRM in skin fibroblasts before (a), during (b), and after (c) the FCCP treatment

its low toxicity, rapid equilibration, and ability to be excited on epifluorescence, confocal, or two-photon microscopes (Fig. 1). In general, two types of approaches can be performed when using TMRM at low concentration (5–25 nM): (1) dynamic measurements to stablish differences in cell–cell signaling or pathophysiological condition, and (2) a comparison of populations of cells that have been previously exposed to different conditions. It is also possible to quantify the change in membrane potential as a function of time in response to a given stimulus. Using TMRM and a confocal microscope, bright pixels can be localized in mitochondria leaving black pixels as a background (Fig. 2). Therefore, when plotting the fluorescence distribution of the pixels along the cell, a high standard deviation will result, and these data can be divided by the mean fluorescence values to calculate changes over a period of time (Fig. 3) [7]. After applying an external variable, illustrated here with the response to an uncoupler, carbonyl cyanide p-trifluoromethoxyphenylhydrazone

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Fig. 3 (a) Mean intensity and its SD over a region of interest, and (b) the ratio of SD/mean fluorescence signal

(FCCP), the fluorescence from mitochondria will decrease and that of the cytosol will increase as the dye redistributes between compartments. Overall, the cell fluorescence will remain stable in the short term, due to the distribution of the dye between the mitochondria and the cytosol. In the longer term, the dye will start to leak from the cell. Thus, we have set up a simple experiment where we can interrogate the response of the mitochondrial membrane potential.

2

Materials Recording medium: Prepare recording medium containing phenol red-free DMEM (Gibco, A1443001) with 10 mM glucose, 1 mM glutamine, 10 mM HEPES, adjusted to pH 7.4. TMRM stock solution: Prepare a 10 mM stock solution of TMRM by dissolving 5 mg of TMRM in 1 ml of dimethyl sulfoxide (DMSO; Sigma-Aldrich, D2650). Make aliquots and store them at 20  C protected from light. Staining solution: Prepare a staining solution of the TMRM reagent by diluting to 25 nM in recording medium. Additionally, use Hoechst 33342 at 5 μM for visualizing the nuclear structure in live cell imaging (see Note 1). Prepare a fresh mixture when working different days and protect the solution from direct exposure to light. Drugs: Prepare 1 μM of FCCP, following the manufacturer’s instructions. This compound will depolarize the mitochondrial

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membrane potential, so then, changes in fluorescence can be compared among conditions over time. This same approach can be used to explore potential mitochondrial toxicity of drugs, or of any biological reagents of interest. Additional reagent: Dulbecco’s Phosphate-Buffered Saline (DPBS; Gibco, 14190094), glass-bottom 24-well plates (Greiner Bio-One, 662892).

3

Methods

3.1 Seeding and Staining Skin Fibroblasts

The entire procedure should be conducted at room temperature unless otherwise stated in the text. 1. Seed cells at appropriate density (1  104/well) in glassbottom 24-well plates or similar imaging plates 2–3 days before imaging (see Note 2). 2. Wash the cells twice with pre-warmed Recording Medium without Hoechst 33342 or DPBS. 3. Incubate the cells with recording medium containing 25 nM of TMRM and 5 μM Hoechst 33342 for 30 min at 37  C. 4. Replace the medium with Recording Medium without Hoechst 33342. 5. Set up the stage and focus. 6. To perform live imaging of cells incubated with TMRM and record dynamic changes, confocal laser scanning microscopy (LSM 880, Carl Zeiss Inc., see Note 3), with the application of live time-series program, is used (see Note 4). Apply low-resolution if needed and attenuated laser power. Excite TMRM with a 561 nm Argon laser with an output power of 0.2 mW. Hoechst 33342 needs to be excited at 405 nm, and it is recommended to start with low laser power when using this channel, as TMRM is very phototoxic (see Note 5). 7. Start acquiring images in basal condition during a few seconds. 8. Add FCCP into the imaging plate, according to previous determined concentration. Wait for the dissipation of the mitochondrial potential and the distribution of the TMRM. 9. Save the time-series and use an appropriate software to obtain the data. Fiji (image J) could be useful for this purpose.

3.2

Data Analysis

1. Define a threshold for the images to exclude black pixels and then select and calculate the average fluorescence intensities from all ROIs of each cell for TMRM and its standard deviation of pixels over the arbitrary threshold.

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2. Select regions next to the cells to calculate the background fluorescence intensity. 3. Subtract the average background fluorescence intensity from average fluorescence intensities of ROIs in each cell for each time point using Microsoft Excel. 4. Plot the standard deviation over the mean signal and analyze the results.

4

Notes 1. Hoechst 33342 concentration can be modified depending on cell type, and it needs to be manipulated with caution since Hoechst strains are known mutagens. Staining intensity may increase with time if samples are imaged without washing. 2. Different cell densities can be used to determine the mitochondrial membrane potential. It is a common practice to plate cells and wait for them to grow until confluence before imaging. 3. A different microscope can be used depending on the availability. 4. Make sure of carefully adding the FCCP. Disturbing the plate will change the focus, and the final analysis cannot be performed. 5. If needed, it is possible to use the same concentration of TMRM (25 nM) and microscope settings to analyze the steady-state measurements of pretreated cells. We recommend acquiring multiple Z-stacks of the cells and use Fiji (image J) to obtain the maximal intensity projection of the images. Then, compare the mean intensities.

Acknowledgments Funding for assay development and manuscript preparation was provided by a scholarship from the National Agency for Research and Development (ANID)/Scholarship Program/DOCTORADO BECAS CHILE/2019—7220052 which supports GEV. References 1. Briston T, Roberts M, Lewis S, Powney B, Staddon JM, Szabadkai G, Duchen MR (2017) Mitochondrial permeability transition pore: sensitivity to opening and mechanistic dependence on substrate availability. Sci Rep 7(1):10492

2. Duchen MR (2004) Mitochondria in health and disease: perspectives on a new mitochondrial biology. Mol Asp Med 25(4):365–451 3. Bazhin AA, Sinisi R, De Marchi U, Hermant A, Sambiagio N, Maric T, Budin G, Goun EA

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(2020) A bioluminescent probe for longitudinal monitoring of mitochondrial membrane potential. Nat Chem Biol 16(12):1385–1393 4. Wacquier B, Combettes L, Dupont G (2020) Dual dynamics of mitochondrial permeability transition pore opening. Sci Rep 10(1):3924 5. Bhosale G, Duchen MR (2019) Investigating the mitochondrial permeability transition pore in disease phenotypes and drug screening. Curr Protoc Pharmacol 85(1):e59

6. Granatiero V, Pacifici M, Raffaello A, De Stefani D, Rizzuto R (2019) Overexpression of mitochondrial calcium uniporter causes neuronal death. Oxidative Med Cell Longev 2019: 1681254 7. Duchen MR, Surin A, Jacobson J (2003) Imaging mitochondrial function in intact cells. Methods Enzymol 361:353–389

Chapter 23 Investigating Mitochondrial Ca2+ Dynamics in Isolated Mitochondria and Intact Cells: Application of Fluorescent Dyes and Genetic Reporters Gauri Bhosale and Michael R. Duchen Abstract Mitochondrial Ca2+ buffering is a hallmark of eukaryotic cellular physiology, contributing to the spatiotemporal shaping of the cytosolic Ca2+ signals and regulation of mitochondrial bioenergetics. Often, this process is altered in a pathological context; therefore, it can be scrutinized experimentally for therapeutic intervention. In this chapter, we describe fluorescence and bioluminescence measurement of mitochondrial Ca2+ in both isolated mitochondria and intact cells. Key words Mitochondrial calcium, Fluorescence calcium imaging, Calcium-sensitive genetic probes, Bioluminescence calcium sensing

1

Introduction Cytosolic Ca2+ signals are shaped by mitochondrial Ca2+ buffering, linking an increase in bioenergetic demand to ATP production. When cytosolic Ca2+ concentrations rise above a certain threshold, energized mitochondria take up Ca2+ in an electrogenic manner, primarily through the MCU complex. The activity of three ratelimiting mitochondrial enzymes in the TCA cycle is in turn upregulated by Ca2+ ions, thus providing more reducing power in the form of NADH to the electron transport chain for ATP synthesis [1]. Mitochondrial Ca2+ levels are further modulated by efflux through the Na+/Ca2+ exchanger at physiological levels, as a pathological increase in mitochondrial Ca2+ levels can induce the opening of the mitochondrial permeability transition pore (mPTP), ultimately resulting in bioenergetic collapse and cell death. Not surprisingly, perturbations in mitochondrial Ca2+ buffering have been widely implicated in neurodegenerative disease (reviewed in [2]). Therefore, monitoring dynamic changes in mitochondrial

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Ca2+ levels can illuminate pathological mechanisms underlying disease etiology. The techniques to monitor mitochondrial Ca2+ are continuously evolving; here, we will briefly describe four different techniques that are routinely used in our laboratory. 1. Use of extramitochondrial Ca2+-sensitive fluorescent dyes with isolated mitochondria in a fluorescence plate reader. 2. Use of intracellular AM-ester-based Ca2+-sensitive cationic fluorescent dyes in intact cells. 3. Use of fluorescence-based genetic probe, 2mtGcamp6, in intact cells: Ideal for ratiometric single-cell analysis to identify relative inter-population and intra-population differences. 4. Use of luminescence-based genetic probe, mitoaequorin, in intact cells: Ideal for cell population based mitochondrial Ca2+ analysis.

2

Materials

2.1 Measuring Mitochondrial Ca2+ in Isolated Mitochondria Using a Fluorescence Plate Reader

1. Isolated mitochondria. 2. Fluo-5N, pentapotassium salt (see Note 1). 3. MSK buffer: 75 mM Mannitol, 25 mM Sucrose, 5 mM Potassium phosphate monobasic, 20 mM Tris–HCl, 100 mM Potassium chloride, 0.1% (w/v) BSA, pH adjusted to 7.4. 4. Succinate. 5. Rotenone. 6. Calcium Chloride. 7. 96-well clear or black plastic plate. 8. Fluorescence plate reader with the following optical requirements: Excitation ¼ 494 nm, Emission ¼ 516 nm, and integrated fluid addition function.

2.2 Measuring Mitochondrial Ca2+ in Intact Cells Using Rhod-2, AM, or 2mtGcaMP6 with Fluorescence Microscopy

1. Rhod-2, AM ester (see Note 1). 2. MitoTracker Green. 3. 2mtGcamp6 adenovirus or plasmid. 4. MitoTracker Deep Red. 5. Recording buffer: Any medium that is compatible with your cells, ideally the medium used to grow the cells without phenol red and with additional pH buffers such as HEPES. 6. Glass coverslips/imaging dishes. 7. For Rhod-2: Fluorescence microscope with the following optical requirements: Excitation ¼ 552 nm, Emission ¼ 560–610 nm.

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For 2mtGcaMP6: Fluorescence microscope with the following optical requirements: Isosbestic Ca2+-independent excitation ¼ 410 nm, Ca2+-dependent excitation ¼ 480 nm, Emission ¼ 500–550 nm. 8. Cytosolic Ca2+ agonist: for example, glutamate or ATP. 9. Ionomycin. 2.3 Measuring Mitochondrial Ca2+ in Intact Cells Using Mitoaequorin Using a Luminescence Plate Reader

1. Mitoaequorin adenovirus or plasmid. 2. Krebs Ringer Buffer: 125 mM NaCl, 5.5 mM D-Glucose, 5 mM KCl, 20 mM HEPES, 1 mM Na3PO4, 1 mM Glutamine, 100 mM Pyruvate, 1.2 mM CaCl2, pH adjusted to 7.4. 3. White 96-well polystyrene plate. 4. Coelenterazine (commercially available). 5. Luminescence plate reader with integrated fluid addition function. 6. Cytosolic Ca2+ agonist: for example, glutamate or ATP. 7. Digitonin.

3

Methods

3.1 Measuring Mitochondrial Ca2+ Uptake in Isolated Mitochondria Using an Extra-Mitochondrial Fluorescence Indicator

1. Add 10 mM succinate and 1 μM rotenone to the MSK buffer; this promotes mitochondrial respiration and therefore allows mitochondrial Ca2+ uptake by maintaining the mitochondrial membrane potential. Prepare a 1 μM Fluo-5N solution using the buffer (Fluo-5N buffer). 2. Make a 100 μM CaCl2 solution in the Fluo-5N buffer, to be added by the integrated fluid injections of the plate reader. 3. Set up the fluorescence plate reader: excitation and emission settings, priming the injector, and setting the temperature to 30  C. Adjust the gain using a well containing the 100 μM CaCl2 as a maximum reading. 4. Make a 0.5 mg/ml mitochondrial suspension in the Fluo-5N buffer and add 100 μl of the suspension per well of the 96-well plate in triplicate for each condition. 5. Add a bolus of 10 μM CaCl2 to each well and measure the response every 10 s. Addition of the CaCl2 will cause an increase in fluorescence intensity, followed by a decay as the Ca2+ is buffered by the mitochondria. Fitting a slope to this decrease can provide a readout for rate of mitochondrial Ca2+ uptake. 6. The protocol can be further modified to include repeated 10 μM CaCl2 additions to determine the calcium retention capacity of mitochondria, beyond which the mitochondria

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undergo permeability transition (mPTP). As a result, all the buffered Ca2+ is released causing a massive increase in fluorescence. The area under the curve can provide a surrogate for the amount of Ca2+ buffered or the “calcium retention capacity,” which can be compared between different conditions. Fluid additions should be set up at regular intervals (see Note 2). 3.2 Measuring Mitochondrial Ca2+ in Intact Cells Using Rhod-2, AM, or 2mtGcaMP6 with Fluorescence Microscopy

1. The seeding density of cells will be determined by cell type, where the user should aim to have a healthy monolayer of cells on the day of imaging, allowing single-cell resolution (see Note 3). 2. For Rhod-2: On the day of the assay, aspirate the growth medium and wash the cells. Replace the medium with recording buffer containing 5 μM Rhod-2, 100 nM MitoTracker Green, and 0.002% pluronic acid. Incubate in the dark at room temperature for 30 min. Aspirate the dye solution, wash the cells, and replace with pre-warmed recording buffer (see Note 4). 3. For 2mtGcaMP6: Transfect the cells with the 2mtGcamp6 with your method of choice, 48 h before imaging. On the day of the assay, aspirate the growth medium, wash the cells, and replace it with pre-warmed recording buffer (see Note 5). 4. If possible, maintain the temperature of the cells at 37  C during the course of the experiment. 5. Set up the microscope with the optical paths described: For

Rhod-2: Excitation Emission ¼ 560–610 nm

¼

552

nm,

For

MitoTracker Green Emission ¼ 500–550 nm

¼

490

nm,

For

2mtGcaMP6: Isosbestic Ca2+-independent excitation ¼ 410 nm; Ca2+-dependent excitation ¼ 480 nm; Emission ¼ 500–550 nm

For

MitoTracker Deep Red Emission ¼ 665–700 nm

¼

644

nm,

6. Adjust the parameters of your fluorescence system for both the optical paths (i.e., exposure/laser power of excitation, master gain, digital offset). If using a confocal, adjust the pinhole to ensure the best compromise between signal:noise ratio and resolution. For 2mtGcaMP6: The isosbestic excitation track will have a lower intensity and provides a measure for expression of the protein. This in turn offers an internal Ca2+-independent control for each cell, allowing relative comparisons between individual cells and populations. For the initial experiments, we recommend confirming mitochondrial localization of the

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genetic probe by co-staining with commercially available live mitochondrial dyes like MitoTracker Deep Red. 7. For measuring basal mitochondrial Ca2+ levels, take multiple images at the desired magnification. We recommend imaging between 10 and 50 cells for relatively homogenous populations. 8. In order to look at kinetic responses to external stimuli, set up a time series with a relatively short time interval (1 s). You might have to reduce resolution to speed up acquisition. 9. After capturing 3–5 baseline images, administer your calcium agonist carefully using a Gilson pipette or perfusion system if available. Please ensure that the z-plane is not disturbed. 10. Monitor the change in fluorescence in both optical tracks. For Rhod-2 and MitoTracker Green: Ideally you should see an increase in intensity in the Rhod-2 image. For 2mtGcaMP6: Ideally you should see an increase in intensity in the 488 nm image, while the 410 nm image remains unchanged. 11. Wait for 3–5 min, for the fluorescence trace to plateau and/or return to baseline. Following this, add the ionomycin solution at a final concentration of 2 μM as a positive control. Ionomycin is an ionophore that should result, causing an influx of Ca2+ into the mitochondria, causing a significant increase in the intensity of the Rhod-2 image and the 488 nm image of 2mtGcaMP6. 12. Once the images have been obtained, they can be analyzed using any image processing software (We recommend ImageJ). Subtract the background from each image. Draw regions of interest around every cell in a field of view. For Rhod-2 and MitoTracker Green: Rhod-2 is nonspecific and will be visible in both the cytosol and the mitochondria. In order to isolate the mitochondrial signal, use the MitoTracker Green image to create a mask (threshold the image and apply a binary mask) and use this mask to measure the mitochondrialspecific Rhod-2 signal. For quantitative comparisons, the Rhod-2 signal can be calibrated using the following equation, where Rf (the dynamic range of the indicator) can be determined using commercially available calibration kits. Fmax is the maximum signal obtained after application of ionomycin [3]      2þ  F 1 F Ca ¼ KD   = 1 F max RF F max For 2mtGcaMP6: Apply a threshold to only include mitochondrial pixels, before measuring mean intensity for both the 488 nm image and the 410 nm image. A ratio of 488 nm/

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410 nm intensities should provide a relative measure of mitochondrial Ca2+ concentrations and can be used to compare across conditions. Exclude cells that appear to be very bright (implying overexpression of the plasmid), as it might provide inaccurate results. 3.3 Measuring Mitochondrial Ca2+ in Intact Cells Using the Bioluminescent Probe, Mitoaequorin

1. Plate 30–50k cells per well in the 96-well plate, in triplicate for each condition. 2. Transduce the cells with the mitoaequorin adenovirus 48 hours prior to the assay (see Note 5). 3. On the day of the assay, aspirate media from wells and replace it with Krebs Ringer buffer containing 5 μM coelenterazine. Incubate the plate in the dark for 2 hours at 37  C to reconstitute the mitoaequorin with its prosthetic group, coelenterazine (see Note 6). 4. While the plate is being incubated, prepare the following solutions: l

l

5 ml of Ca2+ agonist (10 stock) in Krebs ringer buffer (suggested final concentrations: 10 μM glutamate, 100 μM ATP). 5 ml of 1 mM digitonin and 100 mM CaCl2 in H2O.

5. Set up the luminescence plate reader to add the Ca2+ agonist solution after 5 s and the digitonin solution after 30 s. Measure every second for a minute. The digitonin addition provides the luminescence value for aequorin in saturating Ca2+ conditions (see Note 7). 6. Run the assay at 37  C and collect the data for analysis. 7. Accurate mitochondrial [Ca2+] for each timepoint can be calculated as follows: L ¼ ðluminescence ðcounts per secondÞÞ  ðminimum luminescence countÞ L max ¼ ðSum of luminescence counts from the given timepoint till completion of experimentÞ

 Ratio ¼ Ca2þ ðMÞ ¼

L

1=n

L max

ratio þ ðratio K TR Þ  1 K R  ðratio K R Þ

where KR ¼ 10,366,185, KTR ¼ 120, n ¼ 2.99 for wild-type aequorin at 37  C (see Note 8) [4]. The parameters that can be measured in the above techniques include, and are not limited to, rate of mitochondrial Ca2+ uptake, maximum [Ca2+] peak, quantity of Ca2+ buffered, mitochondrial Ca2+ efflux rate to name a few. Resting mitochondrial [Ca2+] can

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also be determined by the fluorescence microscopy techniques, provided appropriate calibrations are carried out (see Note 9).

4

Notes 1. For all methods described, the Kd of the chosen probe or dye should be compatible with the expected changes in [Ca2+], to avoid erroneous results. 2. The number of cycles needs to be determined for your setup. Ideally mitochondrial Ca2+ uptake should be complete, i.e., the fluorescence should return to baseline, before the next addition of Ca2+. 3. For certain cell types like neurons, low seeding density can have a negative impact on viability, whereas for other cell types, higher levels of confluency can cause crowding and cell death. The seeding density therefore needs to be optimized for your experimental setup. 4. The AM ester derivative of Rhod-2 is crucial as it is cell permeable. Furthermore, we have observed better mitochondrial loading of the dye at room temperature, due to decreased activity of the esterases, allowing the dye to cross the mitochondrial membrane before being cleaved and trapped in the cytosol. Pluronic acid is a mild detergent which facilitates dispersal of the AM ester dyes and therefore promotes cell loading. 5. The method of transfection will be dependent on cell type and the available technology. In our laboratory, we routinely use adenovirus to transduce primary cell cultures for higher transfection efficiency, which is essential for the mitoaequorin protocol. 6. If possible, always use Kreb’s Ringer Buffer as coelenterazine is hydrophobic and serum containing medium can reduce bioavailability. If using serum containing medium for sensitive cultures, you might have to use higher concentrations of coelenterazine. 7. Due to the nature of fast kinetics, we recommend running the assay in well mode (i.e., measure the entire time series in one well at a time). Automated fluid additions will further eliminate any loss of data due to delayed readings. 8. The constants for calibrating the aequorin measurements vary based on the type of aequorin and the temperature. These values can be found online.

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9. Mitochondrial Ca2+ uptake or mitochondrial Ca2+ efflux can be pharmacologically targeted to examine the effects on mitochondrial Ca2+ dynamics using these techniques.

Acknowledgments Funding for assay development and manuscript preparation was provided by the BBSRC and Mundipharma. References 1. Denton RM (2009) Regulation of mitochondrial dehydrogenases by calcium ions. Biochim Biophys Acta 1787(11):1309–1316 2. Duchen MR (2012) Mitochondria, calciumdependent neuronal death and neurodegenerative disease. Pflugers Arch 464(1):111–121 3. Maravall M, Mainen ZF, Sabatini BL, Svoboda K (2000) Estimating intracellular calcium

concentrations and buffering without wavelength ratioing. Biophys J 78(5):2655–2667 4. Granatiero V, Patron M, Tosatto A, Merli G, Rizzuto R (2014) The use of aequorin and its variants for Ca2+ measurements. Cold Spring Harb Protoc 2014(1):9–16

Chapter 24 A Plate Reader-Based Measurement of the Cellular ROS Production Using Dihydroethidium and MitoSOX Chih-Yao Chung and Michael R. Duchen Abstract Intracellular reactive oxygen species (ROS) act as an important signaling transductor in cells, regulating almost every aspect of cell biology. Measurements of ROS production thus, offer links between oxidative stress and cell pathophysiology. Here, we describe a simple screening assay in intact adherent cells by fluorescence microplate readers, using dihydroethidium (DHE) and MitoSOX to measure cytosolic superoxide and mitochondrial superoxide production, respectively. This assay enables a quick and reliable assessment of ROS generation in a well-controlled environment. Key words Reactive oxygen species, Dihydroethidium, MitoSOX, Mitochondria, Plate reader

1

Introduction Mitochondria are the main sources of reactive oxygen species (ROS). Intracellular ROS play an important role in cell signaling, regulating cell growth/death, differentiation, and metabolism under pathophysiological conditions. Aberrant ROS production also causes oxidative stress, damaging proteins and nucleic acids (such as DNA). Thus, measurement of the rate of generation of ROS is therefore essential for providing links between mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Here, we describe a simple method using dihydroethidium (DHE) and MitoSOX for measuring cytosolic superoxide and mitochondrial superoxide production, respectively, in intact adherent cells by fluorescence microplate readers. This method is useful for screening experiments and can provide an overview of general ROS production. DHE is an intracellular superoxide indicator which is oxidized by superoxide to forms a red fluorescent product, 2-hydroxyethidium. MitoSOX is a cationic derivative of DHE targeted to the mitochondria and thus used to measure superoxide

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_24, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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production in the mitochondrial matrix, which reacts with superoxide anions in a similar way with DHE. Despite some limitations [1], DHE and MitoSox are still convenient probes for measuring intracellular oxidative stress and redox status.

2

Materials 1. A fluorescent microplate reader with filter sets capable of measuring fluorescence at Excitation (ex) ¼ 480–520 nm, Emission (em) ¼ 570–600 nm. 2. Black, clear-bottom, tissue culture-treated 96-well plates. 3. Recording media: DMEM no phenol red with HEPES is used for cancer cell linens and fibroblasts; DMEM/F12 no phenol red with HEPES is used for neuron-progenitor cells (NPC) and neurons (see Notes 1 and 2). Adjust nutrients, such as glucose, sodium pyruvate, and L-glutamine, according to your culture conditions/experimental settings. 4. Dihydroethidium (DHE): make a 5 mM stock solution in DMSO and aliquot to avoid freeze thaw cycles. Prepare a working solution of 5 μM DHE in recording media by diluting the stock solution (see Note 3). Protect from light. 5. MitoSOX: make a 5 mM stock solution in DMSO and aliquot to avoid freeze–thaw cycles. Prepare a working solution of 5 μM MitoSOX in recording media by diluting the stock solution (see Note 4). Protect from light. 6. Antimycin A (AA): AA is a mitochondrial complex III inhibitor, which causes excessive ROS production when applied. Make a 10 mM stock solution in ethanol and prepare a 150 μM working solution by diluting 15 μl of the stock solution into 1 ml recording media. 7. N-Acetyl Cysteine (NAC): NAC is an antioxidant that raises intracellular glutathione levels by providing cysteine, thereby upregulating the rate-limiting step in the synthesis of the glutathione tripeptide. Freshly make a 300 mM NAC working solution by dissolve 10 mg NAC in 200 μl recording media. Discard after experiments.

3

Methods 1. Two days before the experiments, cells are seeded into the clear-bottom 96-well plates (approximately 10,000 cells per well for cancer cell lines and 30,000–40,000 cells per well for fibroblasts and NPC; see Note 5) and cultured in the media

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suited for users’ cell lines. Ensure that cells reach around 80–90% of confluence on the day of the experiment (see Note 6). 2. Designate at least two wells as positive (AA) and two wells as negative (NAC) controls (see Note 7). 3. Carefully wash the positive and negative control wells with PBS twice and add 150 μl recording media. 4. Add 10 μl NAC to negative control wells and incubate for 30 min at 37  C. 5. After that, add 10 μl AA to positive control wells and incubate for an additional hour at 37  C. 6. Carefully wash all the wells with PBS twice and add 100 μl of 5 μM either DHE or MitoSOX to each well. 7. Set a plate reader to ex ¼ 480–520 nm/em ¼ 570–600 nm (see Note 8) and measure the fluorescence at intervals of 2–5 min for 30–40 min (see Notes 9 and 10). 8. Plot the fluorescence intensity against time points of each well (Fig. 1). The slope of the linear range was used for analysis to determine and represents the rate of ROS production (see Notes 11 and 12) [2].

Fig. 1 A scheme describing how kinetics of ROS production is measured in intact adherent cells, as the rate of increase in red fluorescence intensity over 40 min incubation with DHE/MitoSOX

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Notes 1. Other media without phenol red can also be used if it is more appropriate for specific cell types. 2. HEPES is not necessary if 5% CO2 can be delivered by a plate reader. 3. Working concentration of DHE for measuring ROS production ranges from 0.5 to 5 μM [1–5]. Therefore, conducting a titration for DHE working concentrations to optimize your results is encouraged. The cyto- and mitotoxicity and the redistribution of fluorescence to the nuclei at higher concentrations of these probes should be considered. 4. Working concentration of MitoSOX ranges from 2 to 10 μM [1–4]. Therefore, conducting a titration to optimize your results is encouraged. 5. The initial cell numbers and the duration can and should be adjusted for specific cell types. As cell size and volume vary in cell types, performing a seeding titration is recommended. 6. If the confluence varies among wells with different cell types/ treatments, normalization of the ROS production should be considered. For example, DAPI/Hoechst 33342 staining or BCA assay to obtain the relative cell numbers/amount of protein in each well. 7. Blank wells (without cells) can also be designated to remove the background change in fluorescence intensity caused by spontaneous oxidation of the probe to its fluorescent metabolite [6]. 8. DHE itself (reduced form) displays blue fluorescence with ex/em ¼ ~370/420. Monitoring the blue fluorescence change of DHE simultaneously can be used as an internal control. 9. The interval depends on the measurement speed of a plate reader and the number of wells. 10. Studies suggest that DHE is stable within 30 min [4], and MitoSOX tends to accumulate in the nucleus after approximately 40 min [3]. 11. If blank wells are designated, remove the average fluorescence background from the measurements at each time point. 12. DHE and MitoSOX can also undergo unspecific oxidation into ethidium, which is hard to distinguish from 2-dihydroxyethidium because of fluorescence spectra overlapping. This should be taken into account when interpreting the results [1].

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Acknowledgments Funding for assay development and manuscript preparation was provided by a grant from the Kephalos foundation. References 1. Wojtala A, Bonora M, Malinska D et al (2014) Chapter thirteen - methods to monitor ROS production by fluorescence microscopy and fluorometry. In: Galluzzi L, Kroemer GBT-M (eds) Conceptual background and bioenergetic/mitochondrial aspects of oncometabolism. Academic Press, Boston, pp 243–262 2. Chung C-Y, Singh K, Kotiadis VN, et al (2021) Constitutive activation of the PI3K-AktmTORC1 pathway sustains the m.3243A>G mtDNA mutation. Nat Commun 12:6409 3. Sundqvist M, Christenson K, Bjo¨rnsdottir H et al (2017) Elevated mitochondrial reactive oxygen species and cellular redox imbalance in human NADPH-oxidase-deficient phagocytes. Front Immunol 8:1828

4. Wang Q, Zou M-H (2018) Measurement of reactive oxygen species (ROS) and mitochondrial ROS in AMPK knockout mice blood vessels. Methods Mol Biol 1732:507–517. https:// doi.org/10.1007/978-1-4939-7598-3_32 5. Kro¨ller-Scho¨n S, Daiber A, Schulz E (2018) Modulation of vascular function by AMPK: assessment of NO bioavailability and surrogates of oxidative stress. In: Neumann D, Viollet B (eds) AMPK: methods and protocols. Springer, New York, pp 495–506 6. Tollefson KE, Kroczynski J, Cutaia MV (2003) Time-dependent interactions of oxidantsensitive fluoroprobes with inhibitors of cellular metabolism. Lab Investig 83:367–375. https:// doi.org/10.1097/01.LAB.0000059934. 53602.4F

Chapter 25 Analysis of Organization and Activity of Mitochondrial Respiratory Chain Complexes in Primary Fibroblasts Using Blue Native PAGE Kritarth Singh and Michael R. Duchen Abstract Blue Native polyacrylamide gel electrophoresis (BN-PAGE) is a well-established technique for the isolation and separation of mitochondrial membrane protein complexes in a native conformation with high resolution. In combination with histochemical staining methods, BN-PAGE has been successfully used as clinical diagnostic tool for the detection of oxidative phosphorylation (OXPHOS) defects from small tissue biopsies from patients with primary mitochondrial disease. However, its application to patient-derived primary fibroblasts is difficult due to limited proliferation and high background staining. Here, we describe a rapid and convenient method to analyze the organization and activity of OXPHOS complexes from cultured skin fibroblasts. Key words Oxidative phosphorylation, Supercomplex, In-gel activity, Primary fibroblasts, Mitochondria

1

Introduction BN-PAGE is a convenient and inexpensive separation method that allows the isolation and determination of the oligomeric state of membrane protein complexes in their native state [1]. In combination with histochemical staining and immunodetection, it allows rapid identification of enzymatic and assembly defects in mitochondrial respiratory chain (MRC) complexes, and thus has led to the development of this techniques as a clinical diagnostic tool for studying the OXPHOS defects in patients with monogenic mitochondrial disorders [2, 3]. The functional study of the OXPHOS system using BN-PAGE has provided important insights into the organization of individual complexes within the inner mitochondrial membrane (IMM). In mammals, the four multimeric enzyme complexes of MRC— complex I, II, III, and IV (CI, CII, CIII, and CIV)—are organized

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_25, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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into higher-order structures known as supercomplexes (SCs) or respirasomes [4]. The formation of SCs by different stoichiometric combinations of MRC complexes requires specific assembly factors. For example, Cox7a2l (renamed as supercomplex assembly factor I (SCAFI)) acts as the primary factor for SC formation by mediating the stable interaction between CIII and CIV [5]. Although, it is widely accepted that OXPHOS system is organized into both individual MRC complexes and SCs such that both structures coexist within the IMM, the mechanism of SC formation and its overall role in regulating mitochondrial function are not well understood [6]. Recently, the functional studies of SCs have demonstrated that the supramolecular organization of MRC complexes mediates fine tuning of cell growth in response to different metabolic cues through the formation of different SCs, confers structural stability to CI, limits ROS generation, and facilitates electron channeling [7, 8]. This protocol describes a rapid and convenient method to analyze the organization and activity of SCs from primary fibroblast with high resolution. BN-PAGE has been used to identify OXPHOS defects from tissue biopsies such as liver and skeletal muscle; however, its application to primary fibroblast is limited [3]. Primary fibroblasts derived from skin biopsies, routinely used as in vitro disease model, are characterized by non-exponential growth in culture. Therefore, a minimum cell density of 4.0
106 cells is required to achieve a reasonably pure yield of mitochondrial preparation from cultured cells [9]. Secondly, the resolution of SCs and individual complexes is hindered due to the high background staining which occur as a result of high levels of co-migrating proteins [3]. Here, we provide an optimized protocol for the visualization and analysis of MRC complexes from as low as 5
106 cells with good resolution. The basic principles of the BN-PAGE remain unchanged. Nonionic detergents are used for the solubilization of isolated mitochondrial membrane protein complexes. Mild detergents with low delipidating properties such as digitonin preserve the supramolecular association of MRC complexes. After solubilization of mitochondria and centrifugation, the anionic dye, Coomassie G-250 is added to the supernatant which binds to all membrane proteins owing to its hydrophobic properties. Dye binding imposes a charge shift that causes all proteins to migrate to the anode at pH 7.5 during electrophoresis. The negatively charged protein surfaces repel each other and prevent the formation of membrane protein aggregates. Furthermore, the negatively charged dye-associated membrane proteins become soluble in detergent-free solution which minimizes the risk of denaturation of detergent-sensitive protein complexes. However, the proteins are not resolved based on the charge/mass ratio but according to the pore size of the acrylamide gradient gels which reduces protein migration velocity

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as a function of protein mass and can completely stop protein migration at a mass-specific pore size limit [10]. We simplify our method in three major steps: steps 1–8 describe mitochondria isolation from the primary fibroblasts, steps 9–15 describe solubilization and BN-PAGE run, following that one can opt to perform either the in-gel activity staining for different mitochondrial complexes (steps 16–21) or immunoblotting (steps 22–30).

2

Materials Primary fibroblast culture: Five days before performing the experiment, seed approximately 3
106 cells in culture medium (Dulbecco’s modified Eagle’s medium supplemented with 10% (v/v) fetal bovine serum and penicillin–streptomycin 50 U/ml and 50 μg/ml, respectively) per 150 cm2 tissue culture dish. Replenish the culture medium on the third day (see Note 1). Use Milli-Q-purified water or equivalent in all buffer recipes and protocol steps. Mitochondria isolation buffer (MIB): 1 M sucrose—dissolve 342.3 g of sucrose in 1 l of distilled water; prepare 40 ml aliquots; store at 20  C. 0.1 M Tris/MOPS—dissolve 12.1 g of Tris in 500 ml of distilled water, adjust pH to 7.4 using MOPS powder, bring the solution to 1 l and store at 4  C. 0.1 M EGTA/Tris— dissolve 38.1 g of EGTA in 500 ml of distilled water, adjust pH to 7.4 using Tris powder, bring the solution to 1 l, and store at 4  C. Prepare 25 ml of MIB by adding 2.5 ml of 0.1 M Tris–MOPS and 250 μl of EGTA/Tris to 5 ml of 1 M sucrose. Bring the volume to 25 ml with distilled water. Adjust pH to 7.4. Add 1
protease inhibitor cocktail (Roche, 11697498001) fresh, prior to mitochondrial isolation. Dulbecco’s phosphate-buffered saline (DPBS; Gibco, 14190094), 0.25% Trypsin–EDTA (Thermo Fisher Scientific, 25200056), Pierce BCA Protein Assay Kit (Thermo Scientific, 23225). Gel electrophoresis: 5% Digitonin (Thermo Fisher Scientific, BN2006), 4
NativePAGE™ sample buffer (Thermo Fisher Scientific, BN20032), NativePAGE™ 5% G-250 sample additive (Thermo Fisher Scientific, BN2004), 20
NativePAGE™ running buffer (Thermo Fisher Scientific, BN2001), Native PAGE™ Cathode Buffer Additive (20
), NativePAGE™ Novex 3–12% Bis-Tris protein gels (1.0 mm, 10 well, Thermo Fisher Scientific, BN2011BX10), NativeMark Unstained Protein Standard (Thermo Fisher Scientific, LC0725), Colloidal blue staining kit (Thermo Fisher Scientific, LC6025). Anode buffer–freshly prepare 1000 ml 1
NativePAGE™ Anode Buffer with water.

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Cathode buffer: (1) Dark blue cathode buffer—add 10 ml of NativePAGE™ Cathode Buffer Additive (20
) and 10 ml NativePAGE™ Running Buffer (20
) to 180 ml anode buffer and mix well. (2) Light blue cathode buffer—add 1 ml of NativePAGE™ Cathode Buffer Additive (20
) and 10 ml NativePAGE™ Running Buffer (20
) to 189 ml anode buffer and mix well. Prepare fresh each time for immediate use. Substrates for CI, CII, and CIV activity: For CI activity—freshly prepare 20 ml of 2 mM Tris/HCl buffer (pH 7.4) from 1 M Tris/ HCl containing 2.5 mg/ml Nitrotetrazolium Blue chloride (NBT) and 0.1 mg/ml NADH. For CII—freshly prepare 5 ml of 0.5 mM Tris/HCl buffer (pH 7.4) containing 25 NBT, add 200 μl 1 M sodium succinate, 8 μl 250 mM phenazine methosulfate (prepare 250 mM stock in DMSO) and bring the solution to 10 ml with water. For CIV—freshly prepare 20 ml solution containing 50 mM potassium phosphate (pH 7.4), 5 mg of 3,30 -diaminobenzidine tetrachloride (DAB), and 10 mg of cytochrome c. Immunoblotting: 20
NuPAGE transfer buffer (Thermo Fisher Scientific, NP0006-1)—freshly prepare 200 ml of 1
NuPAGE transfer buffer with 20% (v/v) methanol in water; 10
TrisBuffered Saline (TBS) (Biorad, 1706435)—freshly prepare 1
TBS with 0.1% (v/v) Tween-20 (Sigma, P9416) (TBS-T), 5% (w/v) skim milk (Millipore, 1153630500) in TBS-T, ImmunBlot PVDF membrane (Bio-Rad, 1620177), OxPhos complex kit antibody cocktail (Thermo Fisher Scientific, 45-7999). Prepare 10 ml primary antibody solution (1:1000) in 5% skim milk in TBS-T, Horseradish peroxidase (HRP)–conjugated IgG secondary antibody (Sigma, A9044). Prepare 10 ml secondary antibody solution (1:5000) in 5% skim milk in TBS-T and Chemiluminescent reagent (e.g., Luminata Forte Western HRP substrate, Merck Millipore, WBLUF0100). Additional materials and equipment: 1.5-ml Eppendorf tubes, 1-ml syringe (e.g., BD Luer-Lok. 309628), 25-gauge 1-in. needle (e.g., BD Microlance 3), centrifuge, Protein gel electrophoresis chamber system (XCell SureLock Mini-Cell, Thermo Fisher, EI0001), Power supply (e.g., PowerPac HC High-Current Power Supply, Bio-Rad, 1645052), Mini-gel wet-transfer system (XCell II™ Blot Module, Thermo Fisher, EI9051), Blot imaging system (e.g., ChemiDoc XRS+ System with Image Lab Software, Bio-Rad, 1708265), and ImageJ2 software (National Institutes of Health).

3

Methods

3.1 Isolation of Mitochondria from Primary Fibroblasts

The entire procedure should be conducted on ice, using refrigerated equipment maintained at 4  C.

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1. Remove the medium from the cells and wash the cells once with DPBS and detach the cells using trypsin solution at room temperature (RT). Add DMEM to stop trypsinization and transfer the cell suspension to 15-ml Falcon tube. 2. Centrifuge cells at 600
g for 5 min at 4  C. Discard the supernatant and resuspend the cells in ice-cold DPBS. Centrifuge cell suspension again at 600
g for 5 min at 4  C. 3. Discard the supernatant and resuspend cells in 800 μl of ice-cold MIB and transfer in a 1.5-ml Eppendorf tube. Incubate the cell suspension for 5 min on ice. 4. Homogenize the cells by drawing the solution into a 1-ml syringe fitted with a 25-gauge needle expel it back into the 1.5-ml tube on ice. Take care to expel the solution against the inside wall of the tube as to utilize that force for cell membrane disruption. 5. Repeat the homogenization steps for a total of five times and centrifuge at 600
g for 10 min at 4  C (see Note 2). 6. Transfer the supernatant (700 μl, avoiding the supernatant close to the pellet containing cell and nuclear debris) to a 1.5ml tube and centrifuge at 8000
g for 10 min at 4  C. Discard supernatant and resuspend pellet containing mitochondria in 800 μl cold MIB. Centrifuge again at 8000
g for 10 min at 4  C (see Note 3). 7. Discard the supernatant and resuspend the pellet in 100 μl of cold MIB (see Note 4). 8. Measure mitochondrial protein concentration using the BCA assay and aliquot 100 μg of mitochondrial protein into 1.5-ml tube, centrifuge at 10,000
g for 10 min at 4  C. Discard supernatant and keep the pellet on ice for performing BN-PAGE (see Note 5). 3.2 Solubilization of Mitochondrial Protein and BN-PAGE

9. Solubilize the mitochondrial membrane by adding 20 μl sample buffer cocktail (Table 1) to 100 μg mitochondrial protein. Gently mix the pellet without the formation of bubbles and incubate solubilized mitochondria on ice for 20 min (see Note 6). 10. Centrifuge at 20,000
g for 20 min at 4  C then collect 20 μl of supernatant into a new 1.5-ml tubes. Add 1.5 μl of Coomassie G-250 sample additive to the above supernatant (see Note 7). 11. Set up electrophoresis system by placing the NativePAGE 3–12% gradient gel in the XCell SureLock™ Mini-Cell apparatus (see Note 8).

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Table 1 Preparation of sample buffer cocktail and quantity of detergent and dye required for 100 μg protein Digitonin/ protein ratio Protein (g/g)

4
sample buffer (μl)

5% Final Digitonin Water volume (μl) (μl) (μl)

5% Digitonin/dye Coomassie ratio (g/g) G-250 (μl)

100 μg 6

5

12

8

3

20

1.5

12. Add 50 ml of dark blue cathode buffer in the inner chamber and check for any leakage. Also, wash the wells with 1 ml dark blue cathode buffer. 13. Fill the inner chamber with remaining cathode buffer and gently load 20 μl sample and protein standards (5 μl) into the wells using a P10 tip. Fill the outer chamber with 600 ml running buffer. 14. Set power supply at 100 V and continue electrophoresis for 30 min until the sample has entered the gel. 15. After the blue running front has moved about one-third of the desired total running distance, remove cathode buffer using a 10-ml pipet or suction tube and fill inner chamber with 200 ml light blue cathode buffer. Continue the run at 200 V for 60 min or until the blue running front reaches gel end (see Note 9). 3.3 In-Gel Activity Staining for CI, CII, and CIV

16. Carefully remove gel after completion of the run. If the samples are limiting, the gel can be cut into strips using a Gel Knife and washed with either Tris/HCl or phosphate buffer (pH 7.4) (see Note 10). 17. For CI activity: Incubate the gel strip in 20 ml of complex I substrate solution at RT. Appearance of violet bands within 15–30 min of incubation is indicative of CI activity (Fig. 1a) (see Note 11). 18. For CII activity: Incubate the gel strip in 10 ml complex II substrate solution for 1–2 h at RT. Appearance of violet band is indicative of CII activity (Fig. 2a). 19. For CIV activity: Incubate the gel strip in 10 ml complex IV substrate solution for 1–2 h at RT. Appearance of brown bands is indicative of CIV activity (see Note 12). 20. For CIV + CI activity: Incubate the gel strip first in complex IV substrate solution. After the appearance of appropriate brown signal, wash the gel with Tris/HCl buffer (pH 7.4) and incubate in complex I substrate solution.

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Fig. 1 BN-PAGE, followed by in-gel activity assay to visualize the SCs and individual complexes from isolated mitochondria of primary fibroblasts. 100 μg of mitochondrial protein (6 g/g digitonin/protein ratio and 8 g/g digitonin/dye ratio) was used for BN-PAGE and in-gel activity for mitochondrial complexes were performed. (a) CI activity is shown in violet (20 min incubation). (b) CIV in brown (1 h incubation). (c) CII activity is shown in violet (1 h incubation)

21. Stop reaction with 10% acetic acid when the appropriate signal is observed. Wash the gel with water and document the gel using a scanner. 3.4

Immunoblotting

As mentioned above, if the samples are limiting, the gel can be cut in half after the run and used for in-gel activity and immunoblotting. 22. Activate PVDF membrane with methanol and transfer it to 1
NuPAGE transfer buffer. 23. Set up the blotting apparatus using the XCell II™ Blot Module as per the manufacturer’s instructions. Prepare the gel/membrane assembly and blotting pads in the cathode core. Remove any trapped air bubbles between the gel and membrane ensuring complete contact of all components. 24. Slide the anode core on top of the pads and fill the blot module with transfer buffer until the gel/membrane assembly is covered. 25. Fill the outer chamber with 600 ml of deionized water and transfer for 2 h at 20 V.

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Fig. 2 BN-PAGE, followed by immunoblotting or Coomassie staining to visualize the SCs and individual complexes. 100 μg of mitochondrial protein (6 g/g digitonin/protein ratio and 8 g/g digitonin/dye ratio) was used for BN-PAGE and the expression levels of SCs and individual complexes was detected by either (a) immunoblotting using an OxPhos antibody cocktail or (b) Coomassie staining and destaining

26. Following transfer, incubate the membrane in methanol for 5 min for background destaining (see Note 13). 27. After complete destaining, incubate the membrane in 5% skim milk in TBS-T for 1 h to block the membrane. 28. Incubate membrane with primary antibody solution overnight at 4  C. 29. Wash the blot three times with TBS-T for 5 min each followed by the incubation in primary antibody solution for 1 h at room temperature (see Note 14). 30. Wash the blot again with TBS-T and visualize the expression levels of SCs and individual complexes using a chemiluminescent reagent and a blot imaging system (Fig. 2a) (see Note 15). Alternatively, the expression levels of SCs and individual complexes can also be visualized using the colloidal blue staining kit as per the manufacturer’s instructions (Fig. 2b). The expression levels of individual bands can be determined using densitometry analysis in ImageJ2 software.

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Notes 1. Ensure that cells reach 100% confluency. The total cell number should be at least 5
106 to obtain a minimum yield of 100 μg of mitochondrial pellet. 2. Do not exceed more than five strokes. This would introduce nuclear DNA contamination into the mitochondrial preparation and hinder sample run during BN-PAGE. 3. Minimize the excess pipetting during the transfer of supernatant. Use a wide bore pipette tip (cut tip end with sharp blade) or transfer by tilting to another 1.5-ml tube. 4. Avoid diluting mitochondria with buffer. Mitochondria retain their functionality when stored in a concentrated form. 5. The mitochondrial pellet from step 5 can be stored at 80  C for several months. Divide the mitochondria in appropriate aliquots before storage to avoid the freeze–thaw cycles. 6. The detergent/protein ratio is very critical for solubilizing the mitochondrial membrane proteins. For 100 μg of mitochondrial protein, the digitonin/protein ratio is 6 g/g. The concentration of digitonin should be adjusted according to the protein concentration, which is given by the expression:   DigitoninðgÞ w Digitonin stock ¼6
V1 ¼ v ProteinðgÞ 7. The detergent/dye ratio is also very critical for a good resolution of SCs and individual complexes as well as in reducing the background staining during electrophoresis. The concentration of dye should be adjusted to give a detergent/dye ratio of 8 g/g. 8. The wells should not be left empty as this majorly distorts the gel. Load 20 μl of 1
sample buffer to any unused lanes. 9. For a better separation of the bands, the gel can be run at low power, 150 V for 2 h. 10. To remove the gradient gel using the Gel Knife by handling it from the bottom, which is stronger and therefore chances of breakage are minimized. 11. Avoid the prolonged incubation of gel strip in CI substrate solution as this will increase the background staining. Once the appropriate violet signal starts appearing after 15–30 min of incubation stop the reaction. 12. Due to the common substrates of CIII and CIV, it is difficult to detect in-gel activity of CIII. This protocol does not describe the method for in-gel activity of CIII.

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13. Properly destain the PVDF membrane with methanol as Coomassie dye binds to membranes and thereby reduces protein binding capacity. Do not use nitrocellulose membranes, since these membranes cannot be destained under the conditions described above. 14. Alternatively, the membrane can be reprobed with different antibody for each mitochondrial complexes. 15. Take multiple exposures of the blot to capture faint signal from the SCs as well as the strong signal for individual complexes.

Acknowledgments This work was supported by GOSH/Sparks Charity grant “From the biology of EPG5 to the pathophysiology of Vici syndrome” to M.R.D. References 1. Schagger H, Cramer WA, von Jagow G (1994) Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis. Anal Biochem 217(2):220–230 2. Pitceathly RD, Rahman S, Wedatilake Y, Polke JM, Cirak S, Foley AR, Sailer A, Hurles ME, Stalker J, Hargreaves I, Woodward CE, Sweeney MG, Muntoni F, Houlden H, Taanman JW, Hanna MG, U.K. Consortium (2013) NDUFA4 mutations underlie dysfunction of a cytochrome c oxidase subunit linked to human neurological disease. Cell Rep 3(6): 1795–1805 3. Van Coster R, Smet J, George E, De Meirleir L, Seneca S, Van Hove J, Sebire G, Verhelst H, De Bleecker J, Van Vlem B, Verloo P, Leroy J (2001) Blue native polyacrylamide gel electrophoresis: a powerful tool in diagnosis of oxidative phosphorylation defects. Pediatr Res 50(5):658–665 4. Acin-Perez R, Fernandez-Silva P, Peleato ML, Perez-Martos A, Enriquez JA (2008) Respiratory active mitochondrial supercomplexes. Mol Cell 32(4):529–539 5. Perez-Perez R, Lobo-Jarne T, Milenkovic D, Mourier A, Bratic A, Garcia-Bartolome A, Fernandez-Vizarra E, Cadenas S, Delmiro A, Garcia-Consuegra I, Arenas J, Martin MA,

Larsson NG, Ugalde C (2016) COX7A2L Is a mitochondrial complex III binding protein that stabilizes the III2+IV supercomplex without affecting respirasome formation. Cell Rep 16(9):2387–2398 6. Enriquez JA (2016) Supramolecular organization of respiratory complexes. Annu Rev Physiol 78:533–561 7. Lapuente-Brun E, Moreno-Loshuertos R, Acin-Perez R, Latorre-Pellicer A, Colas C, Balsa E, Perales-Clemente E, Quiros PM, Calvo E, Rodriguez-Hernandez MA, Navas P, Cruz R, Carracedo A, Lopez-Otin C, PerezMartos A, Fernandez-Silva P, FernandezVizarra E, Enriquez JA (2013) Supercomplex assembly determines electron flux in the mitochondrial electron transport chain. Science 340(6140):1567–1570 8. Shoubridge EA (2012) Supersizing the mitochondrial respiratory chain. Cell Metab 15(3): 271–272 9. 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 10. Wittig I, Braun HP, Schagger H (2006) Blue native PAGE. Nat Protoc 1(1):418–428

Chapter 26 Multiplexing Seahorse XFe24 and ImageXpress® Nano Platforms for Comprehensive Evaluation of Mitochondrial Bioenergetic Profile and Neuronal Morphology Smijin K. Soman, Maryann Swain, and Ruben K. Dagda Abstract The measurement of mitochondrial function has become imperative to understand and characterize diseases characterized by bioenergetic alterations. The advancement of automation and application of high-throughput technologies has propelled our understanding of biological complexity and facilitated drug discovery. Seahorse extracellular flux (XFe) technology measures changes in dissolved oxygen and proton concentration in cell culture media, providing kinetic measurements of oxidative phosphorylation and glycolytic metabolism. ImageXpress® Nano is an automated fluorescent microscope with the ability to perform high-content, fast, and robust imaging in multi-well formats. In this chapter, we present a comprehensive protocol to multiplex the Seahorse XFe24 analyzer with the ImageXpress® Nano high content imaging microscope to provide a comprehensive yet rigorous profile of bioenergetics and its correlation to neuronal function and morphology. Key words Bioenergetics, OCR, ECAR, High-throughput imaging, Neuron, Neurodegeneration, Primary neuronal culture

1

Introduction The mitochondrion is the most important intracellular site for essential bioenergetic processes of the cell. Several critical processes, including oxidative phosphorylation, calcium homeostasis, ROS production, fatty acid oxidation, and the Krebs cycle, are facilitated by mitochondria [1–3]. In mammalian cells, mitochondria produce 80% of ATP for endergonic cellular functions through OXPHOS or oxidative phosphorylation; the rest is produced anaerobically, mainly through glycolysis. Any disruption in the bioenergetic pathways is detrimental to cellular function and leads to pathological outcomes, including neurodegeneration [4]. Oxidative phosphorylation is a catabolic process that oxidizes nutrients in order to establish a proton gradient that is coupled to the electron transport

Namrata Tomar (ed.), Mitochondria: Methods and Protocols, Methods in Molecular Biology, vol. 2497, https://doi.org/10.1007/978-1-0716-2309-1_26, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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chain to synthesize ATP [5]. Toxins such as MPTP can inhibit the physiological function of the Complex 1 subunit, disrupting the electron transport chain [6]. Also, loss-of-function mutations in mitochondrial genes or nuclear-encoded mitochondrial proteins can cause mitochondrial dysfunction [7]. It is worth noting that mitochondrial dysfunction is amplified in high energy-consuming cells such as neurons and muscle cells. Mitochondrial dysfunction is a hallmark in the pathophysiology of neurodegenerative disease, including Parkinson’s disease, where reduced expression and activity of the Complex 1 subunit is distinct [8]. It is thereby evident that mitochondrial function is crucial for healthy cells, which underscores the need to develop innovative tools assaying mitochondrial function to better understand the complex interplay between bioenergetic pathways and their relation to neurodegeneration (e.g., loss of dendrites, caspase-mediated cell death, and necrosis). Although mitochondrial function can be measured through several determinants including, mitochondrial membrane potential, ATP turnover, MTT reduction, and mitochondriaderived NADPH levels, measuring oxygen consumption rates (OCRs) using dye-label techniques is the current technique of choice. In this book chapter, a method to assay mitochondrial function using Seahorse XFe24 technology and concurrent imaging of cells has been described. 1.1

Rationale

1.2 Principle and Working

Technology that measures oxygen consumption rates (OCRs) as a proxy of mitochondria function via the XFe24 BioAnalyzer has emerged as a cutting-edge method to measure cellular bioenergetics (OXPHOS and glycolysis). However, a significant limitation of the XFe24 BioAnalyzer is that mitochondrial function data derived from this instrument is not associated with morphological data of cells, which can provide crucial scientific information to understand how extracellular stimuli, toxins, or pharmacologically active compounds affects cell viability and health [9, 10]. Therefore, improvising the robust XFe24 BioAnalyzer technology by combining simultaneous imaging capability to quantify morphology and expression patterns enhances the understanding of bioenergetic pathways in correlation with cellular function. In this book chapter, the hypothesis to be tested is that treatment with exogenous BrainDerived Neurotrophic Factor (BDNF) will enhance bioenergetics and provide neuroprotection by activating the Trkβ receptor and downstream mechanisms in primary cortical neurons leading to increased neuronal connectivity (increased dendrite length/ complexity). XFe24 BioAnalyzer (Agilent Technologies) can measure the flux of oxygen consumption rates (OCRs) and the flux of protons, also known as extracellular acidification rates (ECARs), as a proxy of glycolysis, in the cell medium [11]. OCRs and ECARs are key

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Fig. 1 Basal respiration: OCR induced by basal respiration provides the measure of endogenous ATP demand driven by cellular function and dependent upon ATP utilization, substrate catabolism, and proton leak. Basal respiration rate is an ideal parameter to standardize cell density while performing initial experiments. ATP-linked respiration and proton leak: Oligomycin injection blocks ATP synthase and reduces the OCRs. The remaining rate of mitochondrial respiration, namely proton leak, is due to non-ATP-linked oxygen consumption. Enhanced ATP-linked OCR indicates increased ATP demand and diminished ATP-linked OCR levels, indicating low ATP demand, substrate scarcity, or damage to the oxidative phosphorylation machinery. Enhanced proton leak can be a sign of mitochondrial dysfunction. Maximal respiration: An uncoupler such as FCCP is used to measure maximal respiration. A high FCCP-induced OCR compared with basal OCR indicates that the mitochondria are below the threshold of the maximal rate of electron transport, thereby increasing spare capacity. Oxidative stress-induced mitochondrial dysfunction can reduce spare capacity. Non-mitochondrial oxygen consumption: Non-mitochondrial respiration is due to cellular enzymes (e.g., residing in peroxisomes) and is thereby important to get an accurate measure of mitochondrial respiration in primary neurons

indicators of mitochondrial respiration and glycolysis, respectively (Fig. 1). The assay is performed by insulating a fraction of cell culture media (~7 μl) that is generated when the fluorescent probes are lowered onto the bottom of the well to create a microchamber in the microplate well, which measure the dissolved oxygen (measured as millimeters of mercury pressure per minute) and protons (measured as the change in milli-pH units per minute) in the microchamber per well. OXPHOS consumes oxygen to make water through a four-electron reduction reaction at complex IV. The primary source of proton production in the media of cells grown in culture is lactate generated via glycolysis. Cellular oxygen consumption and proton excretion cause a robust, measurable change to the concentrations of dissolved oxygen and free protons

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in the cell media measured every few seconds by employing solidstate fluorescence-based sensor probes residing 200 μm above the cell monolayer. The chamber is then unsealed (allowing baseline measurements) and re-sealed to repeat the measurement. Experiments providing a metabolic bioenergetic profile are performed by simultaneously measuring OCRs and ECARs under four different conditions (basal, oligomycin, FCCP; carbonyl cyanide p-trifluoromethoxy phenylhydrazone, and rotenone/antimycin A) in each of 20 wells containing cultured cells.

2

Materials

2.1 For Primary Neuronal Culture

Animal

• C57BL/6J timed pregnant (E14) mouse. Reagents

• Dissection media (50 ml)—DMEM media supplemented with FBS: 5 ml (10%), GlutaMAXTM: 500 μl (0.5 mM). • Plating media (50 ml)—Neurobasal media supplemented with FBS: 1 ml (2%), GlutaMAXTM: 125 μl (0.5 mM), B27: 1 ml (2%), Glutamic acid: 106 μl (25 μM), Penicillin/streptomycin: 250 μl (100 U/ml). • Maintenance media (50 ml)—Neurobasal media supplemented GlutaMAXTM: 125 μl (0.5 mM), B27: 1 ml. • Sterile double-distilled water. • Trypan Blue stain. Equipment

• Laminar flow cell culture hood. • Dissecting macroscope with illumination. • Sterile surgical tools. • Sterile Agilent Seahorse XFe24 well Cell Culture Microplate. • Sterile culture dishes (60 mm). • Sterile microcentrifuge tubes (0.5 ml). • Sterile centrifuge tubes (15 ml). • 37
C Incubator with 5% CO2/95% humidity. • Hemocytometer. • 37
C Water bath. 2.2 For Seahorse XFe24

Reagents

• Seahorse XFe24 Calibrant Solution (pH 7.4). • Seahorse XFe24 Extracellular Flux Assay Kit.

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• Seahorse XFe24 Run Medium—XFe24 base medium/Dulbecco’s Modified Eagle’s Medium Base (DMEM) supplemented with 2 mM GlutaMAX™, 1 mM sodium pyruvate, 25 μM DGlucose were used for the Seahorse experiment. An unbuffered medium is essential for measuring changes in pH during glycolysis. • Oligomycin (stock conc. 1 mM). • FCCP (stock conc. 300 μM). • Rotenone (stock conc. 100 μM). • Antimycin A (stock conc. 100 μM). •

D-Glucose.

• Sodium pyruvate. • GlutaMAX™. Equipment

• Agilent Seahorse XFe24 BioAnalyzer. • pH meter. • 37
C Incubator without CO2. • 37
C water-bath. 2.3 For Immunocytochemistry and Nuclear Labeling

Reagents

• Phosphate-Buffered Saline (PBS) (pH 7.4). • Bovine Serum Albumin (BSA). • PBB: 0.5% BSA in PBS. • Fixing solution (4% paraformaldehyde in PBS). • Triton X-100. • DAPI nuclear stain. • Primary antibody (MAP2B). • Alexa Fluor–conjugated secondary antibody.

2.4

For Imaging

2.5 For Operation and Data Analysis

• ImageXpress Nano automated microscope. • Wave software (Agilent Technologies) wave desktop and controller 2.6 software version 2.6.1 released April 2019. • MetaXpress. • GraphPad Prism.

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Methods

3.1 Primary Neuronal Culture 3.1.1 Coating Cell Culture Plates (Day 0)

3.1.2 Culturing Primary Cortical Neurons (Day 1)

Add poly-L-lysine to each well (0.5 ml/well for Agilent Seahorse XFe24 well Cell Culture Microplate) and swirl gently. Incubate the plates at 37
C for 2 h. Aspirate poly-L-lysine using a sterile serological pipette and rinse each well with sterile double-distilled water. Allow plates to air dry inside the laminar flow cell culture hood. Collect acclimatized E15 timed pregnant mouse (C57BL/6J) from the cage and anesthetize the animals using CO2 in a sealed chamber. Wait until the mouse stops moving and fails to respond to the hind-paw pinch before beginning dissection. • Perform cervical dislocation of the mice and place the mouse on a tray for dissection. Open abdominal cavity, remove embryos, and place them in dissection media on ice contained in a 10-ml petri dish. • Embryo micro-dissection: – Transfer embryos to laminar flow cell culture hood having a dissecting microscope with illumination. – Extract embryos from sac and place in a new petri dish with chilled dissection NB/B27 media. – Remove heads from embryos using scissors and place them in a new petri dish with dissection media. – Remove the skin of the head first by carefully peeling it off with fine forceps under the macroscope. Insert the pointed edge of the fine forceps and break the exoskeleton. – Peal the exoskeleton carefully and apply pressure on the sides to push the brain out of the cavity. – Place the whole brain in a new petri dish with dissection media. – Use a syringe and needle (26G) to perform micro-dissection of embryo brain. – Separate the cerebellum from the whole brain. Later, separate the two hemispheres and remove the meninges. Extract cerebral cortex and place it in a 15-ml centrifuge tube on ice. – Dissociate cells by pipetting up and down multiple times (5–7), first using a 1-ml pipette tip, followed by a 200-μl pipette tip. Place the cell suspension on ice. Please note there is no need to use enzymes for these steps as mechanical digestion is gentler to neurons and better for neuronal survival.

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• Place 10 μl of cell suspension in a microcentrifuge tube and add 40 μl Trypan Blue (0.4%). • Add 10 μl of this solution to the hemocytometer. Allow the liquid to disperse through capillary action. • Count cells from multiple grids (4) and calculate cell concentration making sure to exclude cells that are Trypan Blue positive from the overall count: • Cells/ml ¼ (#cells counted/#grids counted) * (dilution factor) * (10,000 cells/ml). • Determine the volume of the cell suspension to be plated: C1*V1 ¼ C2*V2 for cells/well. • Add the determined volume of the cell suspension to plating media (pre-warmed in 37
C water bath). • Add plating media (0.5 ml for 24-well) containing cells to polye L-lysine coated Seahorse XF 24-well plate (make sure to leave wells A1, B3, C4, and D6 empty as background, sensor correction wells) and transfer the plate to a 37
C incubator with 5% CO2/95% humidity.

3.1.4 Media Change (Day 3)

• Check the neurons to see if they are attached, spreading out, and sending out neurites. • Change two/third of the media with maintenance media that does not contain FBS to reduce the possibility of glial overgrowth. Before initiating the Seahorse assay following the treatment with drugs, make sure that the primary neurons are healthy, as noted by their ability to form long dendrites and axons, are not contaminated and are dispersed as monolayers (not clumped). For more information see the notes below (notes section).

3.2 Assessing Bioenergetics in Dissociated Primary Cortical Neurons 3.2.1 Seahorse Extracellular Flux Assay Kit preparation (Hydrate Sensor Cartridge) 3.2.2 Seahorse XFe24 Cell Culture Plate Preparation

• The Seahorse Extracellular Flux Assay Kit comprises a greencolored Seahorse XFe24 sensor cartridge, a transparent 24-well calibration plate, and a pink insert. Take off the sensor cartridge, green lid, and pink insert from the extracellular flux pack kit. Add 0.5 ml of Seahorse XFe24 calibrant per well and reassemble the flux kit. Incubate the flux kit overnight at 37
C non-CO2 incubator.

• Warm the Seahorse XFe24 base media to 37
C. • In the meantime, turn on the Seahorse XFe24 BioAnalyzer and the interface (monitor screen attached to the instrument). Open the Seahorse Wave (version 6 or higher) software and load the protocol.

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• Create protocol: Set temperature at 37
C. – Basal 3 cycles: mix 3 min, wait 2 min, measure 3 min. – Port D drug addition: measurement 15 cycles, mix 3 min, wait 2 min, measure 3 min. – Oligomycin 3 cycles (Port A): mix 3 min wait 2 min measure 3 min. – FCCP 3 cycles (Port B): mix 3 min, wait 2 min, measure 3 min. – Rotenone/antimycin (Port C) 3 cycles: mix 3 min, wait 2 min, measure 3 min. – Assay complete eject plate, view results, save. • Warm up the XFe24 Seahorse BioAnalyzer to 37
C (5 h minimum). • Take the Seahorse XFe24 plate from the 37
C incubator with 5% CO2/95% humidity and place it in the laminar flow cell culture hood. • Using an aspirator pipette, remove approximately 150 μl of the maintenance media and wash the wells (including correction wells) with 1 ml of Seahorse XFe24 run media. Remove all but 150 μl of media from the wells, making sure to aspirate from the side of the wells (or using a 1 mL pipette for more gentle removal) and not directly into the cells. • Add 525 μl of Seahorse XFe24 run media (including correction wells) and keep the plate at 37
C in a non-CO2 incubator for 1 h. 3.2.3 Drug Preparation

While the primary cortical neurons cultured on the 24-well Seahorse culture plate are incubating in the 37
C in a non-CO2 incubator for 1 h, prepare the drugs for injection heat to 37
C. • Oligomycin 10 solution (25 μM): Add 50 μl of 1 mM stock solution of oligomycin to 2 ml of Seahorse XFe24 run media to make a sub-stock of oligomycin (25 μM). Add 56 μl of this sub-stock solution to port A of the calibrated sensor cartridge for a final concentration of 2.5 μM Oligomycin. • FCCP 10 solution (10 μM): Add 66.66 μl of 300 μM stock solution of FCCP to 2 ml of Seahorse XFe24 run media to make a sub-stock of FCCP (10 μM). Add 62 μl of this sub-stock solution to port B of the calibrated sensor cartridge for a final concentration of 1 μM FCCP. • Rotenone/Antimycin A 10 solution (5 μM): Add 100 μl of 100 μM stock solution of rotenone plus 100 μl of 100 μM stock solution Antimycin A to 2 ml of Seahorse XFe24 run media to make a sub-stock of rotenone/Antimycin A (5 μM). Add 69 μl of this sub-stock solution to port C of the calibrated sensor

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cartridge for a final concentration of 0.5 μM Rotenone/ Antimycin A. • BDNF (Experimental drug): Add 6 μl of BDNF (5 ng/μl) into seahorse media for a sum of 75 μl port D of the calibrated sensor cartridge for a final concentration of 30 ng/well. • Ana12 (Experimental drug, BDNF receptor antagonist): Add 2 μl of 61 μM stock (200 nM/well) with a sum of Ana12 and seahorse media 75 μl for injections to port D. • Control all wells filled with 75 μl seahorse media port D. 3.2.4 Bioenergetic Measurement

• Login to the Seahorse Wave software on the interface and set up an experimental template by using the Assay Wizard option. • Group definitions, plate map, protocol, run assay: input the information regarding type of media used (NB/B27), type of cells, drugs employed for the assay (rotenone/antimycin A cocktail, FCCP, oligomycin, experimental drug), and their concentrations. Save the template and be sure that the “Save Directory” and “Save Name” fields contain the relevant experimental information. • Remove the Seahorse Extracellular Flux Assay Kit from the 37
C non-CO2 incubator. Add the drugs to the respective ports in the calibrated sensor cartridge using the concentrations indicated above. • Take the Seahorse Extracellular Flux Assay Kit containing the XFe calibrant in the calibrant plate wells and take it to the warmed-up (37
C) XFe24 Seahorse Analyzer. • Remove the pink insert and the transparent lid of the Seahorse Extracellular Flux Assay Kit and place the sensor cartridge onto the calibrant plate containing the 0.5 ml of XFe calibrant solution per well. • Start the calibration step by clicking on the “Start” button. When the loading door opens, place the sensor cartridge with the calibration plate on the tray. • When the calibration/quality control process is complete, replace the calibration plate with the experimental cell plate and run assay.

3.3 Immunocytochemistry and Nuclear Labeling

• At the end of the mitochondrial stress assay (approximately 3.5–4 h) remove the medium, follow with 1 wash of 1 PBS, making sure to aspirate the media on the side (not directly) of the wells to avoid lifting the delicate monolayer of primary neurons. Make sure the cells are immediately fixed, preferably within 10 min after the assay is completed (see notes below for more information).

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• Fix sample in 4% paraformaldehyde for at least 15 min at room temperature. • Wash monolayer on each well three times in PBS (5 min/wash) to remove any residual PFA. • Permeabilize cells with 0.3% Triton X-100 made in PBS solution for a minimum of 15 min at room temperature. • Wash the cell monolayer on each well one time with PBS. • Wash the cell monolayer five times with PBB (5 min/wash) and block with 2% BSA for 45 min. • Wash the cell monolayer three times with PBB (5 min/wash). • Primary antibody: To stain for dendrites in primary neurons, dilute the desired concentration of rabbit anti-mouse MAP2B in PBB (vortex gently, spin down for 5 min at 10–12K RPM to get rid of any aggregates). • Incubate overnight at 4
C with a gentle motion. • Remove primary antibody from the wells (or save it by placing it back onto the original antibody working stock) and wash each well five times with PBB. • Add secondary antibody (647 anti-rabbit 1:500) into the wells and incubate for 60 min at room temperature. • Wash the cell monolayer five times on each well with PBB. • Counterstain with DAPI (1:100,000) for 30 s in PBS. • Wash the monolayer one time on each well with PBS. • Store the cells in PBS with 0.05% sodium azide to avoid bacterial growth. Prepare for automated fluorescent imaging analyzed for neuronal morphology (analysis of neurite length/ complexity and cell count) as described below. 3.4 Imaging with ImageXpress Nano

• Switch on the ImageXpress Nano microscope, light source, and computer. • Open MetaXpress® software and click acquisition setup. • Place the Seahorse XF24 Cell Culture Microplate in the imaging chamber and close the lid. • Under the configure tab, select 40 objective. Adjust the correction collar if necessary. • Under the plate tab, click the drop-down list and select the Seahorse XFe24 plate. The dimension of the plate based on vendor specifications should be annotated and saved prior to imaging cells for neurite analysis. Select the site to visit tab and select an appropriate number of sites.

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• Under the acquisition tab, select autofocus tab, and enable laser-based autofocusing. In well to well autofocus set focus to well bottom. • Under wavelength, select number, and choose FITC (mouse anti-human MAP2B) (488 anti-mouse) and DAPI filter. Select FITC wavelength and adjust settings. • In the plate map, select a well (right-click the well). • For adjusting image acquisition, select illumination setting, and click focus. Make sure the image exposure is appropriate. Calculate offset to perform automatic focus determination. Click focus again and check post-laser offset, and the image should be in focus. • Repeat for each subsequent wavelength. • Under the run tab, click experiment details and save protocol. • Click acquire plate to begin acquiring plate. 3.5

Data Analysis

3.5.1 OCR and ECAR Determination

• Open the Wave software, select results and scan to the project of interest, and open. • With primary cortical neurons (post-mitotic), normalizing the OCRs using cell counts is appropriate and a valid way as opposed to normalizing to protein concentration. Select the normalization icon input DAPI numbers on the plate map with normalization unit: cell count and a scale factor: 1. This will update all the data and modify all the graphs, calculations, and analyses. • Once the data is normalized, export the data. Select export, and export the seahorse XFe Cell Mito Stress Test Report Generator for generating the results. • Results: summary printout > bar charts > measures sheet > assay parameters per well > sample calculations > and project information. Figure 2 shows an example of enhancement of oxidative phosphorylation (OCR line graph) and of glycolysis (ECAR line graph) in response to exposure of primary cortical neurons exposed to BDNF but not in the presence of ANA12, an antagonist of the TrKB receptor (Fig. 2). • Export GraphPad Prism for performing statistics to run ordinary one-way ANOVA, column analyses, one-way ANOVA, multiple comparisons: compare the mean of each column with the mean of every other column, options: Tukey’s test.

3.5.2 Neurite Length Measurement

The Neurite Outgrowth application module in MetaXpress software simultaneously identifies and measures cell bodies and neurites.

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Fig. 2 (a) Treatment with BDNF (30 ng, 2 h) significantly increases OCRs in primary cortical neurons when compared to control, also, treatment with Trkβ (BDNF) receptor antagonist Ana12 (200 nM, 2 h) significantly decreases OCRs in primary cortical neurons when compared to control. Statistics: Tukey’s multiple comparisons test: Ana12 ( control) vs. BDNF (exp), significant adjusted P value ****