Mitochondrial Bioenergetics (Methods in Molecular Biology, v810)

METHODS

IN

MOLECULAR BIOLOGY™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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Mitochondrial Bioenergetics Methods and Protocols Edited by

Carlos M. Palmeira MitoLab, Department of Life Sciences, Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal

António J. Moreno Department of Life Sciences and IMAR, University of Coimbra, Coimbra, Portugal

Editors Carlos M. Palmeira MitoLab Department of Life Sciences Center for Neurosciences and Cell Biology University of Coimbra Coimbra, Portugal [email protected]

António J. Moreno Department of Life Sciences and IMAR University of Coimbra Coimbra, Portugal [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-381-3 e-ISBN 978-1-61779-382-0 DOI 10.1007/978-1-61779-382-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011938460 © Springer Science+Business Media, LLC 2012 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface The purpose of this book is to provide a definitive treatise on the conceptual principles and practical considerations regarding research initiatives in mitochondrial function. There has been a robust emergence of interest in mitochondrial bioenergetics of late, much of it driven by realization of the impact of drug and environmental chemical-induced disturbances of mitochondrial function as well as hereditary deficiencies and the progressive deterioration of bioenergetic performance with age. These initiatives have fostered the investment of enormous resources into the investigation of genetic and environmental influences on bioenergetics, demanding a certain degree of understanding of the fundamental principles and a level of proficiency in the practice of mitochondrial bioenergetic research. This book is intended as a bench reference for a broad scope of readers, spanning students of mitochondrial bioenergetics to practitioners in the industries of pharmaceutical and environmental sciences as well as mitochondrial genetics. Each section is prefaced with a short treatise introducing the fundamental principles for that section followed by chapters describing the practical principles and assays designed to derive quantitative assessment of each set of parameters that reflect different aspects of mitochondrial bioenergetics. The hope is that this text will help elevate the quality and rate of investigative discoveries regarding disease states associated with environmental or genetic influences on mitochondrial bioenergetics. Coimbra, Portugal Coimbra, Portugal

Carlos M. Palmeira António J. Moreno

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

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1 Overview of Mitochondrial Bioenergetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Vitor M.C. Madeira 2 Evaluation of Respiration with Clark Type Electrode in Isolated Mitochondria and Permeabilized Animal Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Ana M. Silva and Paulo J. Oliveira 3 High-Resolution Respirometry: OXPHOS Protocols for Human Cells and Permeabilized Fibers from Small Biopsies of Human Muscle . . . . . . . . . . . . . . 25 Dominik Pesta and Erich Gnaiger 4 High-Throughput Analysis of Mitochondrial Oxygen Consumption . . . . . . . . . . . . 59 James Hynes, Rachel L. Swiss, and Yvonne Will 5 Modulation of Cellular Respiration by Endogenously Produced Nitric Oxide in Rat Hippocampal Slices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Ana Ledo, R.M. Barbosa, and J. Laranjinha 6 Mitochondrial Membrane Potential (DY) Fluctuations Associated with the Metabolic States of Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Carlos M. Palmeira and Anabela P. Rolo 7 Safranine as a Fluorescent Probe for the Evaluation of Mitochondrial Membrane Potential in Isolated Organelles and Permeabilized Cells . . . . . . . . . . . . 103 Tiago R. Figueira, Daniela R. Melo, Aníbal E. Vercesi, and Roger F. Castilho 8 Fluorescence Measurement of Mitochondrial Membrane Potential Changes in Cultured Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 David G. Nicholls 9 Phenomenological Kinetic and Control Analysis of Oxidative Phosphorylation in Isolated Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Vilmante Borutaite and Rasa Baniene 10 Expression of Uncoupling Proteins in a Mammalian Cell Culture System (HEK293) and Assessment of Their Protein Function. . . . . . . . . . . . . . . . . . . . . . . 153 Martin Jastroch 11 Measurement of Proton Leak and Electron Leak in Isolated Mitochondria . . . . . . . 165 Charles Affourtit, Casey L. Quinlan, and Martin D. Brand 12 Relation Between Mitochondrial Membrane Potential and ROS Formation . . . . . . 183 Jan M. Suski, Magdalena Lebiedzinska, Massimo Bonora, Paolo Pinton, Jerzy Duszynski, and Mariusz R. Wieckowski 13 Use of a Calcium-Sensitive Electrode for Studies on Mitochondrial Calcium Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 António J.M. Moreno and Joaquim A. Vicente

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14 Imaging Mitochondrial Calcium Signalling with Fluorescent Probes and Single or Two Photon Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . Sean M. Davidson and Michael R. Duchen 15 Mitochondrial Permeability Transition Pore and Calcium Handling . . . . . . . . . . . . Renee Wong, Charles Steenbergen, and Elizabeth Murphy 16 Imaging of Mitochondrial pH Using SNARF-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . Venkat K. Ramshesh and John J. Lemasters 17 Redox Equivalents and Mitochondrial Bioenergetics . . . . . . . . . . . . . . . . . . . . . . . . James R. Roede, Young-Mi Go, and Dean P. Jones 18 NMR Methodologies for Studying Mitochondrial Bioenergetics . . . . . . . . . . . . . . . Tiago C. Alves, Ivana Jarak, and Rui A. Carvalho 19 Computational Modeling of Mitochondrial Function . . . . . . . . . . . . . . . . . . . . . . . Sonia Cortassa and Miguel A. Aon Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors CHARLES A FFOURTIT • Buck Institute for Research on Aging, 8001 Redwood Blvd.,Novato, CA 94945-1400, USA TIAGO C. A LVES • Faculty of Sciences and Technology, Department of Life Sciences, University of Coimbra, R. Larga 6, 3030 Coimbra, Portugal M IGUEL A. AON • Johns Hopkins University, School of Medicine, 1059 Ross Bldg., 720 Rutland Ave., Baltimore, MD 21205, USA R ASA BANIENE • Institute for Biomedical Research, Lithuanian University of Health Sciences, Eiveniu str. 4, LT-50009, Kaunas, Lithuania RUI M. BARBOSA • Faculty of Pharmacy, Center for Neurosciences and Cell Biology, University of Coimbra, R. Larga, 3004-504, 3030, Coimbra, Portugal M ASSIMO BONOR A • Department of Experimental and Diagnostic Medicine, Section of General Pathology, Interdisciplinary Center for the Study of Inflammation (ICSI), BioPharmaNet, University of Ferrara, Ferrara, Italy VILMANTE BORUTAITE • Institute for Biomedical Research, Lithuanian University of Health Sciences, Eiveniu str. 4, LT-50009 Kaunas, Lithuania M ARTIN D. BR AND • Buck Institute for Research on Aging, 8001 Redwood Blvd.,Novato, CA 94945-1400, USA RUI A. CARVALHO • Faculty of Sciences and Technology, Department of Life Sciences, University of Coimbra, R. Larga 6, Coimbra 3030, Portugal ROGER FRIGÉRIO CASTILHO • Faculdade de Ciências Médicas, Departamento de Patologia Clínica, Universidade Estadual de Campinas, Campinas, SP 13083-887, Brazil SONIA CORTASSA • School of Medicine, Johns Hopkins University, 1059 Ross Bldg., 720 Rutland Ave., Baltimore, MD, 21205, USA SEAN M. DAVIDSON • Department of Medicine, The Hatter Cardiovascular Institute, University College London, 67 Chenies Mews, London, WC1E 6HX, London, UK M ICHAEL R. DUCHEN • Department of Cell and Developmental Biology and Consortium for Mitochondrial Research, University College London, Gower Street, London WC1E 6BT, UK JERZY DUSZYNSKI • Nencki Institute of Experimental Biology, Polish Academy of Sciences, 02-093,Warsaw, Poland TIAGO R EZENDE FIGUEIR A • Faculdade de Ciências Médicas, Departamento de Patologia Clínica, Universidade Estadual de Campinas, Campinas, SP 13083-887, Brazil ERICH GNAIGER • D.Swarovski Research Laboratory, Department of General and Transplant Surgery, Medical University of Innsbruck, Innrain 52, Christoph-Probst-Platz 6020, Innsbruck, Austria YOUNG-MI GO • Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine, Emory University, 201 Dowman Drive, Atlanta, GA 30322, USA ix

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JAMES HYNES • Luxcel Biosciences, BioInnovation Centre, UCC, Cork, Ireland IVANA JARAK • Center for Neurosciences and Cell Biology, University of Coimbra, R. Larga, 3004-504, Coimbra 3030, Portugal M ARTIN JASTROCH • Buck Institute for Age Research, 8001 Redwood Blvd., Novato 94945, CA, USA DEAN P. JONES • Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine, Emory University, 201 Dowman Drive, Atlanta, GA 30322, USA JOÃO L AR ANJINHA • Faculty of Pharmacy, Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal M AGDALENA LEBIEDZINSK A • Nencki Institute of Experimental Biology, Polish Academy of Sciences, 02-093 Warsaw, Poland A NA LEDO • Center for Neuroscience and Cell Biology, University of Coimbra, R. Larga 6, Coimbra 3004-504, Portugal JOHN J. LEMASTERS • Center for Cell Death, Injury and Regeneration, Medical University of South Carolina, Charleston, SC 29425, USA Departments of Pharmaceutical & Biomedical Sciences, Medical University of South Carolina, Charleston, SC 29425, USA Deparments of Biochemistry & Molecular Biology, Medical University of South Carolina, Charleston, SC 29425, USA Hollings Cancer Center, Medical University of South Carolina, Charleston, SC 29425, USA VITOR M. C. M ADEIR A • Department of Life Sciences, University of Coimbra, R. Larga 6, Coimbra 3030, Portugal DANIELA RODRIGUES M ELO • Faculdade de Ciências Médicas, Departamento de Patologia Clínica, Universidade Estadual de Campinas, Rua Dr Quirino, 1856 – Centro, Campinas, SP 13083-887, Brazil A NTÓNIO J. M. MORENO • Department of Life Sciences, IMAR, University of Coimbra, Apartado 3046, 3001-401 Coimbra, Portugal ELIZABETH MURPHY • Cardiac Physiology Section, Systems Biology Center, NHLBI, NIH, Bethesda, MD 20892, USA DAVID G. NICHOLLS • Buck Institute for Research on Aging, 8001 Redwood Blvd., Novato, CA 94945-1400, USA PAULO J. OLIVEIR A • Center for Neuroscience and Cell Biology, University of Coimbra, R. Larga 6, 3004-517, Coimbra, Portugal CARLOS M. PALMEIR A • MitoLab, Department of Life Sciences, Center for Neurosciences and Cell Biology, University of Coimbra, Apartado 3046, Coimbra 3001-401, Portugal DOMINIK PESTA • D.Swarovski Research Laboratory, Department of General and Transplant Surgery, Medical University of Innsbruck, Innrain 52, Christoph-Probst-Platz 6020, Innsbruck, Austria PAOLO PINTON • Department of Experimental and Diagnostic Medicine, Section of General Pathology, Interdisciplinary Center for the Study of Inflammation (ICSI), BioPharmaNet, University of Ferrara, Ferrara, Italy CASEY QUINLAN • Buck Institute for Research on Aging, 8001 Redwood Blvd., Novato, CA 94945-1400, USA

Contributors

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VENK AT K. R AMSHESH • Center for Cell Death, Injury and Regeneration, Medical University of South Carolina, 86 Jonathan Lucas Street, PO Box 250955, Charleston, SC 29425, USA Departments of Pharmaceutical & Biomedical Sciences, Medical University of South Carolina, 86 Jonthan Lucas Street, PO Box 250955, Charleston, SC 29425, USA Deparments of Biochemistry & Molecular Biology, Medical University of South Carolina, 86 Jonathan Lucas Street, PO Box 250955, Charleston, SC 29425, USA Hollings Cancer Center, Medical University of South Carolina, 86 Jonathan Lucas Street, PO Box 250955, Charleston, SC 29425, USA JAMES R. ROEDE • Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine, Emory University, 201 Dowman Drive, Atlanta, GA 30322, USA A NABELA P. ROLO • Center for Neurosciences and Cell Biology, University of Coimbra, R. Larga 6, Coimbra 3030, Portugal A NA M. SILVA • Center for Neurosciences and Cell Biology, University of Coimbra, R. Larga 6, Coimbra 3030, Portugal CHARLES STEENBERGEN • Department of Pathology, Johns Hopkins Medical School, 600 N. Wolfe Street, Baltimore, MD 20892, USA JAN M. SUSKI, M.SC. • Nencki Institute of Experimental Biology, Polish Academy of Sciences, Warsaw, Poland Department of Experimental and Diagnostic Medicine, Section of General Pathology, Interdisciplinary Center for the Study of Inflammation (ICSI), BioPharmaNet, University of Ferrara, Via Savonarola, 9, Ferrara, Italy R ACHEL SWISS • Cell Based Assays and Mitochondrial Biology, Compound Safety Prediction – WWMC, Pfizer R&D, Eastern Point Rd, Groton, CT 0634, USA A NIBAL EUGÊNIO VERCESI • Faculdade de Ciências Médicas, Departamento de Patologia Clínica, Universidade Estadual de Campinas, 177 Rua Roxo Moreira, Campinas, SP 13083-887, Brazil JOAQUIM A. VICENTE • Department of Life Sciences, IMAR, University of Coimbra, Coimbra, Portugal M ARIUSZ R. WIECKOWSKI • Nencki Institute of Experimental Biology, Polish Academy of Sciences, 02-093,Warsaw, Poland YVONNE WILL • Cell Based Assays and Mitochondrial Biology, Compound Safety Prediction – WWMC, Pfizer R&D, 558 Eastern Point Rd, Groton, CT 06340, USA R ENEE WONG • Cardiac Physiology Section, Systems Biology Center, NHLBI, NIH, Bethesda, MD 20892, USA

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Chapter 1 Overview of Mitochondrial Bioenergetics Vitor M.C. Madeira Abstract Bioenergetic Science started in seventh century with the pioneer works by Joseph Priestley and Antoine Lavoisier on photosynthesis and respiration, respectively. New developments were implemented by Pasteur in 1860s with the description of fermentations associated to microorganisms, further documented by Buchner brothers who discovered that fermentations also occurred in cell extracts in the absence of living cells. In the beginning of twentieth century, Harden and Young demonstrated that orthophosphate and other heat-resistant compounds (cozymase), later identified as NAD, ADP, and metal ions, were mandatory in the fermentation of glucose. The full glycolysis pathway has been detailed in 1940s with the contributions of Embden, Meyeroff, Parnas, Warburg, among others. Studies on the citric acid cycle started in 1910 (Thunberg) and were elucidated by Krebs et al. in the 1940s. Mitochondrial bioenergetics gained emphasis in the late 1940s and 1950s with the works of Lenhinger, Racker, Chance, Boyer, Ernster, and Slater, among others. The prevalent “chemical coupling hypo thesis” of energy conservation in oxidative phosphorylation was challenged and replaced by the “chemiosmotic hypothesis” originally formulated in 1960s by Mitchell and later substantiated and extended to energy conservation in bacteria and chloroplasts, besides mitochondria, with clear-cut identification of molecular proton pumps. After identification of most reactive mechanisms, emphasis has been directed to structure resolution of molecular complex clusters, e.g., cytochrome c oxidase, complex III, complex II, ATP synthase, photosystem I, photosynthetic water splitting center, and energy collecting antennæ of several photosynthetic systems. Modern trends concern to the reactivity of radical and other active species in association with bioenergetic activities. A promising trend concentrates on the cell redox status quantified in terms of redox potentials. In spite of significant development and advances of bioenergetic knowledge, major issues remain mainly related with poor experimental designs not representative of the real native cell conditions. Therefore, a major effort has to be implemented regarding direct observations in situ. Key words: ATP syntase, Bioenergetics, Chloroplast, Chemical coupling hypothesis, Chemiosmotic hypothesis, Citric acid cycle, Complexes II and III, Cytochrome c oxidase, Energy collecting antennæ, Fermentation, Glycolysis, Mitochondria, Photosystem, Radical, Redox potential, Water splitting center

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_1, © Springer Science+Business Media, LLC 2012

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Very often, Bioenergetics is wrongly considered a modern science of the twentieth century. Rather , the pioneers started already in the middle of seventh century, with the works of Joseph Priestley, in London, and Antoine de Lavoisier, in Paris, in parallel with the development of Chemistry. Joseph Priestley (1, 2) described several important gases, including oxygen, named dephlogisticated air. He discovered that plants, in the presence of daylight, could purify the bad air (unsuitable for respiration) resulted from combustions and metal calcinations (oxidations). plants purified air Bad air daylight combustions good for respiration calcinations In Priestley’s terminology, the bad or phlogisticated air (loaded with phlogist, the principle of fire and calcinations) (cf. Phlogiston Theory) turned into dephlogisticated air. Later, Priestley concluded that dephlogisticated air was in fact a fraction of the total air. Antoine de Lavoisier clearly established that Priestley’s dephlogisticated air was an independent gas, initially named vital air and later oxygine (acid generator, in Greek), the acidifying principle of common acids (3, 4). Additionally, Lavoisier carried out precise experiments on animal and human respiration, showing that pure air turned into acide crayeux (lime acid, carbon dioxide) in lungs during respiration. He further demonstrated that there is a constant relationship between the heat released and the amount of air entered in lungs (4). Therefore, the first measurement of a metabolic rate has been reported by Lavoisier ca. 1780. In 1860, Louis Pasteur described fermentation of sugar into alcohol strictly linked to living yeast cells (5). By the time, it was believed that the ferments (enzymes) could only be active inside the living cell “organized” by the “vital principle” (Vitalism Theory). However, by the end of nineteenth century, the chemists Hans Buchner and Eduard Buchner (6, 7) clearly showed that filtered yeast juice (cells absent) was very active in the fermentation of sugar into alcohol, meaning that the ferments are active when “disorganized”, i.e., without the need of any “vital principle”. This basic idea opened the way of modern Biochemistry, allowing detailed studies of metabolic pathways carried out later in twentieth century. Details on fermentation were provided in 1905 by Harden and Young who demonstrated that yeast extracts rapidly fermented glucose into alcohol and that orthophosphate is required and consumed in the phosphorylation of hexose (8, 9). They separated the yeast extract in two types of compounds: zymase, non dialyzable and easily inactivated by heat; cozymase, a dialyzable fraction resistant to heat. The zymase fraction contained the glycolytic enzymes and cozymase, besides orthophosphate, contained NAD+, ADP, and metal ions.

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Full glycolysis pathway has been detailed in 1940s with the major contributions of Embden, Meyeroff, Parnas, Warburg, and other researchers (10). Studies on citric acid cycle started in 1910 with the work of Thunberg, followed by important achievements of Szent-Györgyi in 1935 (5). However, full elucidation has been provided by Krebs (11) who established the condensation reaction as the cycle closing step. Mitochondrial bioenergetics gained emphasis on late 1940s and 1950s with the relevant work of eminent scientists, e.g., Lehninger, Racker, Chance, Boyer, Ernster, and Slater (12). It has been shown that oxidations of NADH, CoQH2, formed at expense of oxidation of citric acid cycle intermediates, resulted in energy yielding effective in ATP synthesis. The coupling of oxidations and ATP synthesis has been explained on the basis of the “chemical coupling hypothesis” (5) proposing that energy of oxidations could be conserved in “high energy compounds” e.g., phosphate esters as in the case of “substrate level phosphorylations” occurring in glycolysis and the citric acid cycle (5). However, these compounds were never demonstrated or isolated. In the 1960s, Mitchell in his “chemiosmotic hypothesis” proposed that the energy of oxidations is transduced into physicochemical states through the establishment of a transmembrane proton gradient generating a transmembrane electric potential and/or a pH gradient (13) described by the famous equation of the “protonmotive force”: Dp = DY - Z DpH

(Z = 59 at 25°C),

Where the transmembrane potential ΔΨ = Ψin − Ψout and the transmembrane ΔpH = pHin − pHout. The protonmotive force (Δp) is expressed in millivolts and represents the electrochemical potential (free energy change) of the transmembrane proton electrochemical gradient divided by the Faraday’s constant. It is analogous to the electromotive force in electricity. This hypothesis is generally accepted and explains conveniently the energy transduction in mitochondria, bacteria, and chloroplasts (14). Δp is provided by protonmotive force generators at the expense of transmembrane proton transport: from matrix to cytoplasm in mitochondria; from cytoplasm to the intermembrane space in bacteria; from stroma to thylakoid space in chloroplasts (14). Protonmotive force development has been detected in all known membrane redox systems (13), viz. mitochondria, bacteria, chloroplasts, and Archaea (15). Protonmotive force generators function in the basis of molecular proton pumps and the ubiquinone pool associated with the

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activity of mitochondrial complex III or equivalent in bacteria and chloroplasts as proposed originally by Mitchell in his looping mechanisms (14). Molecular proton pumps have been identified in mitochondria (16) and bacterial cytochrome c oxidases (17). Proton pumps have been also putatively assigned to complex I (18). The protonmotive force energy may be used in several activities: ATP synthesis, ion transport, orthophosphate transport, nucleotide exchange, transhydrogenase, heat generation, and flagellar motion (10, 14). Redox processes are carried out in complex protein clusters: I (NADH-CoQ oxidoreductase), II (succinate-CoQ oxidoreductase), III (CoQH2-cytochrome c oxidoreductase), and IV (cythochrome c-O2 oxidoreductase) or the equivalents in bacteria. In chloroplasts, complexes I, II, and IV are absent and photosynthetic PSI and PSII clusters are present. Complexes diffuse laterally in the lipid membrane (19) with relatively low diffusion constants for the big complexes (I, II, III, and IV) and the transfer of reducing equivalents is achieved randomly during effective encounters. These are significantly facilitated and accelerated by the ubiquinone pool (associating complexes I and II with III) and cytochrome c (associating complexes III and IV), owing to the fast diffusion constants (19). This strategy involving big complex clusters and small fast components (ubiquinone and cytochrome c or equivalents) is a common motif in all known redox systems: mitochondria, bacteria, and chloroplasts. In several complexes (I and II), redox sequences occur in two distinct segments: a two-electron event (e.g., oxidation of NADH to NAD+ and succinate to fumarate) is followed by an one-electron event (iron-sulfur centers of complexes I and II). The two events are coupled by a flavin center able to process either two or one electron at a time, undergoing intermediate semiquinone species (10). This strategy is a common motif in all redox systems where a redox segment of two-electrons is followed by an oneelectron segment, e.g., bacteria (20), photosynthetic bacteria (20) (synthesis of NADH), chloroplasts (10) (synthesis of NADPH), methanogens (20), and nitrogenase systems in Rhizobium (21). After identification of basic reactive mechanisms, emphasis has been directed to the structure resolution of complex clusters. Detailed structures of cytochrome c oxidases (22, 23), complex III (24), and complex II (25) have been described. Complex I has been also partially resolved (18). Structure of ATP synthase (26) and its clustering with the orthophosphate carrier and the adenine nucleotide carrier (27) (ATP synthasome) have been established. Available also are the structures of photosystem I (28, 29), water splitting center (30), and energy-collecting antennae (31–33) of several photosynthetic systems. The structural trend is still in progress for other bioenergetic assemblies.

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Modern research trends are concerned with oxidative processes related with the formation and reactivity of radical species (oxygen related and other) in association with mitochondrial and other cell activities. A promising trend, attempting a quantitative description, regards the integration of the radical chemistry with the cell redox status in terms of NAD+/NADH, NADP+/NADPH, GSSG/GSH, and other redox balances in relation to redox potentials (34). These efforts may effectively contribute to a clear quantitative appraisal how the redox balances affect the radical chemistry and its involvement in major cell functions, viz. mitochondrial transition permeability, apoptosis, necrosis, enzyme, and gene activities. In spite of the relevant progress in bioenergetic knowledge, there are still major issues and challenges to be accomplished. Most issues relate to poor experimental conditions that roughly deviate from the native cell conditions. Most available data has been collected with isolated preparations which contain fragments of the original chondriom framework. Therefore, it should be not surprising that relevant functions got lost and other severely modified during the crude isolation procedures. Hence, drawn conclusions must be evaluated with caution. Furthermore, most experimental setups and reaction media are diverse from in situ conditions, e.g., temperature is generally set at 25°C (for technical reasons), instead the actual cell temperature (37°C), in experiments of oxygen tracing by electrometry. The chemical composition of media (sucrose, salt concentrations, buffers, pH) often does not minimally relate with cytoplasm condition. Furthermore, oxygen experiments are carried out at saturation (240 μM, at 25°C), a situation far from the expected low oxygen activity in the living cell. Therefore, data on oxygen radicals may be severely questioned. It is of general concern that a significant effort has to be implemented regarding direct observations in situ, looking at bioenergetic activities directly in living cells. This is a tremendous and difficult challenge for the near future. If not accomplished, we will never be certain if the observations are facts or artifacts. References 1. Priestley J (1775) An account of further discoveries in air. Philos Trasact 65:384–394 2. Priestley J (1775) Experiments and observations on different kinds of air, 2nd edn. J Johnson, London 3. Lavoisier AL (1789) Traité elementaire de chimie. Cuchet, Paris 4. Lavoisier AL (1864) In oeuvres de Lavoisier, Tome II. memoires de chimie et de physique. Imprimerie Imperiale, Paris 5. Lehninger AL (1975) Biochemistry. Worth Publishers, New York

6. Buchner E (1897) Alkoholische gärung ohne hefezellen. Berichte der Deutschen Chemischen Gesellshaft 30:117–124 7. Buchner E, Rapp R (1899) Alkoholische gärung ohne hefezellen. Berichte der Deutschen Chemischen Gesellshaft 32:2086–2094 8. Mahler HR, Cordes EH (1971) Biological chemistry, 2nd edn. Harper and Row, New York, p. 495 9. Harden, A. and Young, W. J. 1908. The alcoholic ferment of yeastjuice. Part III. The function of phosphates in the fermentation of

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cytochrome c oxidase from Paracoccus denitrificans. Nature 376:660–669 Zang Z, Huang L, Shulmeister VM et al (1998) Electron transfer by domain movement in cytochrome bc1. Natura 392:677–684 Yankovskaya V, Horsefield R, Törnroth S et al (2003) Architecture of succinate dehydrogenase and reactive oxygen species generation. Science 299:700–704 Abrahams JP, Leslie AGW, Luter R, Walker JE (1994) Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370:621–628 Chen C, Ko Y, Delannoy M, Ludtke J, Chiu W, Pedersen PL (2004) Mitochondrial ATP synthasome. J Biol Chem 23:31761–31768 Deisenhofer J, Epp O, Sinning I, Michel H (1995) Crystallographic refinement at 2.3 Å resolution and refined model of the photosynthetic reaction centre from Rhodopseudomonas viridis. J Mol Biol 246:429–457 Jordan P, Fromme P, Witt HT, Klukas O, Saenger W, Krauss N (2001) Three-dimensional structure of cyanobacterial photosystem I at 2.5 Å resolution. Nature 411:909–917 Ferreira KN, Iverson TM, Maghlaoui K, Barber J, Iwata S (2004) Architecture of the photosynthetic oxygen-evolving center. Science 303: 1831–1838 McDermott G, Prince SM, Freer AA, Hawthornthwaite-Lawless, Papiz MZ, Cogdell RJ, Isaacs NW (1995) Crystal structure of an integral membrane light-harvesting complex from photosynthetic bacteria. Nature 374:517–521 Karrasch S, Bullough PA, Gosh R (1995) The 8.5 Å projection map of the light-harvesting complex I from Rhodospirillum rubrum reveals a ring composed of 16 subunits. EMBO J 14:631–638 Liu Z, Yan H, Wang K, Zhang J, Gui L, An X, Chang W (2004) Crystal structure of spinach major light-harvesting complex at 2.72 Å resolution. Nature 428:287–292 Jones DP (2006) Disruption of mitochondrial redox circuitry in oxidative stress. Chem Biol Interact 163:38–53

Chapter 2 Evaluation of Respiration with Clark Type Electrode in Isolated Mitochondria and Permeabilized Animal Cells Ana M. Silva and Paulo J. Oliveira Abstract In many studies, the evaluation of mitochondrial function is critical to understand how disease conditions or xenobiotics alter mitochondrial function. One of the classic end-points that can be assessed is oxygen consumption, which can be performed under controlled, yet artificial conditions. Oxygen is the terminal acceptor in the mitochondrial respiratory chain, namely at an enzyme called cytochrome oxidase, which produces water in the process and pumps protons from the matrix to the intermembrane space. Several techniques are available to measure oxygen consumption, including polarography with oxygen electrodes or fluorescent/luminescent probes. The present chapter will deal with the determination of mitochondrial oxygen consumption by means of the Clark-type electrode, which has been widely used in the literature and that still remains to be the most reliable technique. We focus our technical description in the measurement of oxygen consumption by isolated mitochondrial fractions and by permeabilized cells. Key words: Mitochondria, Permeabilized cells, Clark type electrode, Cellular respiration, ADP/O ratio, Respiratory control ratio

1. Introduction Aerobic eukaryotic cells present very developed intracellular machinery aimed at energy production, the mitochondrion, where O2 is the terminal electron acceptor of a series of electron transfers through proteins comprising the so-called respiratory chain (1). The diagram in Fig. 1 represents the mitochondrial respiratory chain, where electrons flow from NADH (oxidized in complex I) or succinate (complex II) to O2. The different complexes contain several co-factors and prosthetic groups. Complex I contains FMN and 22–24 iron–sulfur (Fe–S) proteins in 5–7 clusters. Complex II contains FAD, 7–8 Fe–S proteins in three clusters and cytochrome b560.

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_2, © Springer Science+Business Media, LLC 2012

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Fig. 1. Mitochondrial respiratory chain showing the process of oxidative phosphorylation.

Complex III contains cytochrome b (the ultimate electron acceptor in this complex), cytochrome c1, and one Fe–S protein. Complex IV contains cytochrome a, cytochrome a 3, and two copper ions. It is proposed that when two electrons pass through complex I, four protons (H+) are pumped into the intramembrane space of the mitochondrion. In the same manner, as each pair of electrons flows through complex III, four protons are pumped into the intramembrane space. Electrons are then used to reduce O2 to H2O, on cytochrome c oxidase. The protons in intramembrane space return to the mitochondrial matrix down their concentration gradient by passing through the ATP synthase, also called complex V, which synthesizes ATP at the expense of the transmembrane electric potential generated by the proton imbalance between the two sides of the membrane (2). The process of ATP synthesis by mitochondria is termed oxidative phosphorylation (OXPHOS), which describes the coupling between substrate oxidation and ADP phosphorylation. When OXPHOS and electron transfer system (ETS) in particular, needs to be evaluated, an approach to monitor oxygen dynamics is a possibility, especially when coupled to proper substrates and inhibitors of the electron transfer and ADP phosphorylation process (see Note 1).

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Historically, the electrochemical reduction of oxygen was discovered by Heinrich Danneel and Walter Nernst in 1897 (revised by Severinghaus and Astrup (3)). In the 1940s, the technology was tested in biological tissues to follow oxygen supply or availability by the implantation of platinum electrodes (4), in an amperometric approach. The major problem of the direct contact of electrodes with biological preparations was contamination, diminishing the electrochemical response; one example was the contamination of the platinum electrode by compounds such as cyanide and iodide (5). The fundamental innovation introduced by L. Clark (3) was the presence of a permeable membrane, which allows the transport of ions but restricts the contact of higher molecular weight substances (e.g. proteins) with the electrode surface (6). Nevertheless, it is important to know the properties of the chosen oxygen permeable membrane, paying special attention to the electrochemical delays, and the possibility of organic degradation or physico–chemical interactions with some inhibitors such as antimycin A or oligomycin, among others (5), that could still be present from one experiment to the next. It is thus necessary to obtain information from the current concentration relationship, thus maintaining a desired selectivity. The chemical transformations that take place at the electrodes after the passage of electrical current are ruled by Faraday’s law and by the current–voltage equation. In vivo cell respiration, considering organ and whole body organization, is regulated by intracellular non-saturating ADP levels during normal states of activity (7) and increases when submitted to a specific stimulus. One reason to use mitochondria in intact or permeabilized cells is that mitochondria interact with the cytoskeleton network, being grouped in functional clusters in close contact with other cell organelles and structures, probably an essential feature for the correct function of the whole mitochondrial bioenergetic apparatus (8). Cells permeabilization may be established by different methodologies, but the most popular is the one that considers the presence of amphipathic substances. Detergents (such as digitonin and saponin) have been used for selective membrane permeabilization, but their usage conditions must be carefully controlled. By treating cells with specific detergents, it is possible to induce membrane permeabilization by promoting loss of lipid content, although without losing important intracellular proteins; subsequently it will be easier to follow the oxygen consumption in the presence of different substrates, nucleotides, and respiratory inhibitors. Cell permeabilization by electroporation is also technically possible. Electrically permeabilized cells were used to allow the uptake of exogenously added ATP, which is essential for receptormediated activation of the respiratory burst. For example, blood cells as neutrophils, are ATP-depleted prior to permeabilization (9).

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Differences between intact cell respiration and substrate oxidation in digitonin-permeabilized cells can be polarographically measured with a Clark oxygen electrode in a micro-jacketed chamber (10, 11), although quenched-fluorescence oxygen sensing can be also used in intact cells (12). In the next sections of the present book chapter, some representative procedures to prepare permeabilized cell and isolate mitochondria are demonstrated, as well some hypothetical schemes for the assessment of oxygen respiration.

2. Materials 2.1. Cell Culture/ Preparation and Permeabilization 2.1.1. Electropermeabilized Cells: Neutrophils (Adapted from ref. 9)

1. Ficoll 400 and dextran T-500 (Pharmacia LKB Biotechnology Inc.). 2. RPMI-1640, AMP–PNP, ATP, EGTA, GDP, GTP, NADPH, glucose, 2-deoxy-d-glucose, Coomassie Blue, analytical grade salts (Sigma). 3. Bicarbonate-free medium RPMI-1640 (buffered to 7.3, with 25 mM NaHEPES). Sterilize by autoclaving. 4. Permeabilization medium: 140 mM KCl, 1 mM MgCl2, 1 mM EGTA, 10  mM KHEPES (pH 7.0), and CaCl2 enough to reach a final concentration of 100 nM, 1 mM ATP (except for experiments without ATP), and 10 mM glucose (except for ATPdepletion experiments, where is replaced by 2-deoxy-d-glucose). Store it on ice (±4°C). 5. Reaction medium for oxygen consumption assays: 140  mM NaCl, 5 mM KCl, 10 mM HEPES, 1 mM MgCl2, 1 mM CaCl2, 1 mM MgCl2, and 10 mM NaHEPES (pH 7.3). 6. HEPES-RPMI-1640 medium and all cell buffers must be at 37°C previously to usage. 7. All reagents (salts, sucrose, etc.) with analytical grade, from Sigma (St. Louis, MO).

2.1.2. SaponinPermeabilization: Saponin-Skinned Muscle Fibers (Adapted from ref. 8)

1. Isolation/resuspension medium: 10 mM Ca–EGTA (0.1 mM free Ca2+), 20 mM imidazole, 20 mM taurine, 50 mM K-MES, 0.5  mM DTT (dithiothreitol), 6.56  mM MgCl2, 5.77  mM ATP, and 15 mM phosphocreatine (pH 7.1). Store on ice. 2. Saponin (Sigma S-2149) and other reagents (salts, sucrose, etc.) with analytical grade, from Sigma (St. Louis, MO).

2.1.3. DigitoninPermeabilization: Lymphocytes (Adapted from ref. 11)

1. Histopaque 10771 and other reagents (salts, etc.) with analytical grade, from Sigma (St. Louis, MO). 2. Phosphate buffer: 137 mM NaCl, 2 mM KCl, 1 mM KH2PO4, and 10 mM Na2HPO4 (pH 7.4).

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3. Digitonin (Sigma-Aldrich D-141) and other reagents (salts, sucrose, etc.) with analytical grade, from Sigma (St. Louis, MO). 2.2. Isolation of Mitochondrial Fraction

1. Brain isolation buffer (BIB): 225  mM mannitol, 75  mM sucrose, 5 mM HEPES, 1 mM EGTA, and 1 mg/mL bovine serum albumin (fatty acid free) (pH 7.4). 2. Brain resuspension buffer (BRB): 225 mM mannitol, 75 mM sucrose, and 5 mM HEPES (pH 7.4). 3. Liver and heart isolation buffer (LIB and HIB): 250  mM sucrose, 10 mM HEPES, 0.5 mM EGTA, and for liver isolation supplement medium with 1  mg/mL bovine serum albumin (fatty acid free) (pH 7.4). 4. Liver and heart resuspension buffer (LRB and HRB): 250 mM sucrose and 10 mM HEPES (pH 7.4). 5. Protease (subtilisin fraction VIII) and other reagents (salts, sucrose, etc.) with analytical grade, from Sigma (St. Louis, MO).

2.3. Oxygen Electrode 2.3.1. Oxygen Electrodes and Polarographic Systems

Commercially available, being the most common, the following: Oroboros Oxygraph (Physica respirometer, Paar Physica, Oroboros, Austria); Hansatech Instruments Limited (Norfolk, UK); Yellow Springs Instrument Company (YSI Inc., Ohio, USA, the one we use in our laboratory); Gilson Medical Electronics (Middleton, WI); Mitocell micro respirometer (Strathkelvin Instruments Limited, Scotland). See specifications with manufactures.

2.3.2. Oxygen Permeable Membranes

We considered preferentially PTFE films [poly(tetrafluoroethylene), e.g. Teflon] with the reference YSI 5776, Oxygen Probe Service Kit (Yellow Springs, OH, USA).

2.4. Oxygen Consumption Assays: Permeabilized Animal Cells

1. Neutrophils respiration medium: 140 mM NaCl, 5 mM KCl, 10 mM HEPES, 1 mM MgCl2, 1 mM CaCl2, 1 mM MgCl2, and 10 mM NaHEPES (pH 7.3). 2. Saponin-skinned muscle fibers respiration medium: 110  mM mannitol, 60  mM KCl, 10  mM KH2PO4, 5  mM MgCl2, 60 mM Tris–HCl, and 0.5 mM Na2EDTA (pH 7.4). 3. Lymphocytes respiration medium: 300 mM mannitol, 10 mM KCl, 10  mM HEPES, 5  mM MgCl2, and 1  mg/mL bovine serum albumin (fatty acid free) (pH 7.4).

2.5. Oxygen Consumption Assays: Isolated Mitochondria

1. Brain mitochondria respiration medium: 100  mM sucrose, 100  mM KCl, 2  mM KH2PO4, 5  mM HEPES, and 10  mM EGTA (pH 7.4). 2. Heart mitochondria respiration medium: 50  mM sucrose, 100 mM KCl, 1 mM KH2PO4, 10 mM Tris, and 10 mM EGTA (pH 7.4).

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3. Liver mitochondria respiration medium: 135  mM sucrose, 65  mM KCl, 5  mM KH2PO4, 5  mM HEPES, and 2.5  mM MgCl2 (pH 7.4). It must be stressed that other reaction media are possible, using different compositions and if, required, with different osmolarities, although typically media with 250–300 mOsm are used.

3. Methods 3.1. Cell Culture and Permeabilization 3.1.1. Electropermeabilized Cells: Neutrophils (Adapted from ref. 9)

1. Isolate neutrophils from fresh heparinized human blood by dextran sedimentation followed by Ficoll–Hypaque gradient centrifugation. 2. Add NH4Cl for red cell lysis, centrifuge again, and wash with HEPES-buffered RPMI. 3. Count cells (using ZM Coulter Counter) and resuspend them at 107 cell/mL in HEPES-buffered RPMI [for ATP-depleted cells, cell (107/mL) incubation at 37°C, during 5 min, in the specific buffer (with 2-deoxy-d-glucose, instead of glucose), prior to permeabilization]. 4. Electroporation: Sediment cells (with ATP or ATP-depleted) by centrifugation and resuspend in ice-cold permeabilization solution at 107 cells/mL. Add 800 mL cell suspension/cuvette in a Bio-Rad Pulser (although others can be used) and apply two electric discharges (pulses – charges of 5 kV/cm from a 25-mF capacitor) as described by the manufacturer. Sediment cells by centrifugation in 1.5 mL conic tubes (Eppendorf 5415 microcentrifuge), between pulses, and resuspend in ice-cold permeabilization solution (at 107 cells/mL). 5. Electroporated cells are ready to be used by picking cells to a final density of 2 × 106 cell/mL or can be stored on ice up to 15 min.

3.1.2. SaponinPermeabilization: Saponin-Skinned Muscle Fibers (Adapted from ref. 8)

1. Muscle fibers should undergo mechanical separation. Under a microscope, dissect small bundle fibers (submerged on isolation/ resuspension medium ice-cold) at one end and separate down to the muscle mid belly (using fine jewellers forceps). 2. Place muscle bundles in ice-cold isolation medium (8–15 wet weight/mL), add 50 mg/mL saponin, and gently mix the suspension during 30 min, at 4°C. 3. Wash muscle fibers with resuspension medium to remove all saponin. 4. Measure protein by Lowry et  al. (13) method, using a 5-mL aliquot. Consider 0.5 mg/mL for oxygen consumption assays.

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5. Store on ice (muscle fibers in resuspension medium) until start respiration analysis in specific reaction medium. 3.1.3. DigitoninPermeabilization: Lymphocytes (Adapted from ref. 11)

1. Isolate human lymphocytes introducing 5 mL heparinized blood in a tube filled with a solution containing polysucrose and sodium diatrizoate, adjusted to a density of 1.077 g/mL (HISTOPAQUE® 10771-Sigma) (the proportion between Histopaque and blood should be 1/3 and 2/3, respectively). 2. Centrifuge (Eppendorf 5702 centrifuge) at 500 × g for 30 min (20°C) until mononuclear cells form a distinct layer at the plasma-HISTOPAQUE® interface (the blood must remain on top, do not mix). Wash with phosphate-buffered saline (PBS), centrifuge, and resuspend. Repeat the procedure twice. 3. Resuspend in a final volume of 25  mL of the same buffer. Measure protein by the Lowry method (13), using a 5-mL aliquot. Consider 0.5 mg/mL for oxygen consumption assays. 4. Digitonin (1%) is added to lymphocytes just after introduction in the oxygen consumption chamber.

3.2. Isolation of Mitochondrial Fraction

Independently of cell mitochondrial origin, the first important procedure to achieve good and feasible results on oxygen respiration is the isolation of tightly coupled mitochondrial fractions. Several factors have to be taken into account, including the composition and temperature of all the solutions used.

3.3. Mitochondrial Isolation from Brain, Heart, and Liver (from Wistar Rats)

1. After being anesthetized using different possible processes, rats should be sacrificed accordingly with the ethical proceedings established by the “Guide for the Care and Use of Laboratory Animals”, published by the US National Institutes of Health (NIH Publication No. 85–23, revised in 1996).

3.3.1. Brain Mitochondria (Adapted from ref. 14)

2. Decapitate rat (reserve body for other organ extraction) and remove rapidly the cerebellum. Wash and mince it (using sharp scissors) and later homogenize (in a small glass Potter-Elvehjem) at 4°C in 10  mL of brain isolation buffer (BIB) containing 5 mg of bacterial protease. 3. Brain homogenate is brought to 30 mL, and centrifuge 2,000 × g for 3 min (Sorvall RC6 plus centrifuge; SS-34 rotor). 4. Resuspend pellet, including the fluffy synaptosomal layer, in 10  mL BIB containing 0.02% digitonin and centrifuge at 12,000 × g for 8 min. 5. Resuspend brown mitochondrial pellet without the synaptosomal layer in 10  mL BRB and centrifuge at 12,000 × g for 10 min. 6. Resuspend mitochondrial pellet in 300 mL BRB. Store mitochondria on ice until assays start.

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7. Determine mitochondrial protein by biuret method (20) calibrated with bovine serum albumin (BSA). Consider 0.8 mg/mL for oxygen consumption assays, although other protein concentrations can be used. 3.3.2. Heart Mitochondria (Adapted from ref. 15)

1. Open rat chest with scissors and remove the heart. 2. Place heart in isolation buffer (HIB) and cut slowly the organ in little pieces with cold scissors. 3. Wash heart pieces as many times as needed with HIB to remove all blood. 4. Homogenize heart pieces in a Potter-Elvehjem with 20 mL HIB complemented with 35 mL protease (subtilisin fraction VIII or Nagarse). Be careful to maintain temperature under 4°C. 5. The homogenization takes place with 3 or 4 homogenizations with a refrigerated piston (preferably). The suspension is then left under incubation during 1 min on ice. After that period of time, homogenize 2–3 more times. 6. Place heart homogenate in centrifuge tubes and fill up with HIB. 7. The tubes are centrifuged at 12,000 × g during 10 min (Sorvall RC6 plus centrifuge; SS-34 rotor). 8. The supernatant is discharged and the pellet is freed after addition of HIB. Transfer to a smaller glass manual Potter-Elvehjem homogenizer, and promote gentle homogenization. 9. Centrifuge at 2,000 × g during 10  min. Pour supernatant to new refrigerated centrifuge tubes, and fill up with HIB. 10. Centrifuge supernatant at 12,000 × g during 10 min. 11. Discharge the new supernatant, and gently homogenise pellet with a smooth paintbrush, adding heart resuspension buffer (HRB). Place homogenate in centrifuge tubes and fill up with HRB. 12. Centrifuge at 12,000 × g during 10  min. Discharge supernatant and resuspend isolated mitochondria in 200–300  mL in HRB. Store mitochondria on ice until start oxygen respiration assays. 13. Determine mitochondrial protein by biuret method (20) calibrated with BSA. Consider 0.5 mg/mL for oxygen consumption assays, although different mitochondrial protein concentrations can be used, according to the respiratory activity of the preparation.

3.3.3. Liver Mitochondria (Adapted from ref. 4)

1. Open rat abdominal cavity with a scissor and remove liver. 2. Place liver in isolation buffer (LIB) and cut slowly in little pieces with a scissor. 3. Wash liver pieces as many times as needed with LIB to remove all blood.

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4. Homogenize liver pieces in a Potter-Elvehjem with 60  mL LIB. Be careful to maintain temperature under 4°C. 5. Place liver homogenate in centrifuge tubes and fill up with LIB. 6. The tubes are centrifuged at 2,000 × g during 10 min (Sorvall RC6 plus centrifuge; SS-34 rotor). Pour supernatant to new refrigerated centrifuge tubes, and fill up with LIB. Avoid contamination with debris from pellet. 7. Centrifuge supernatant at 12,000 × g during 10 min. 8. Discharge the new supernatant, and gently homogenize pellet with a smooth paintbrush, adding liver resuspension buffer (LRB). Place homogenate in centrifuge tubes and fill up with HRB. 9. Repeat steps 7 and 8. 10. Centrifuge at 12,000 × g during 10 min. Discharge supernatant and resuspend isolated mitochondria in 200–300  mL in LRB. Store mitochondria on ice until start oxygen respiration assays. 11. Determine mitochondrial protein by biuret method (20) calibrated with BSA. Consider 1.5 mg/mL for oxygen consumption assays, although alternatives in mitochondrial protein content can be considered. 3.4. Oxygen Electrode Preparation and Maintenance

The Clark type electrode is, as it is expected from the name, an electrode that measures oxygen on a catalytic platinum surface. The classical Clark type is basically composed of two electrodes (cathode and anode). The electrical signal arises from the current developed on the cathode. The voltage supply unit (associated with a galvanometer) feeds the system with electrons, and when oxygen diffuses through an oxygen permeable membrane (commonly Teflon-based) to the platinum electrode, the cathode is reduced. The current is proportional to the oxygen tension in the solution (16) (see Note 2) where mitochondria or cells are placed. The silver ions combine with chloride ions in solution, precipitating silver chloride over the silver electrode, accordingly with the overall equation 4Ag(s) + 4Cl− + O2 + 4H+ + 4e− → 4AgCl×2H2O (17, 18). The electric signal can be analogically recorded using a flatbed recorder, or the electrode output current can pass through an analogto-digital converter and later be analysed through a signal transducer coupled to a personnel computer (as exemplified in ref. 19) (Fig. 2).

3.5. Respiratory Parameters

Two important criteria used to assess the integrity or quality of a determined mitochondrial fraction after isolation or after incubation with test compounds are the respiratory control ratio (RCR) and the ADP/O. The RCR is a measure of the coupling between substrate oxidation and phosphorylation and basically informs the researcher

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Fig. 2.  General representation of the Clark-type electrode and experimental setup. (a) The biological preparations (mitochondrial or cell suspensions) are introduced in a well-defined volume of media (depending of the preparations), inside a thermostated incubation chamber (1) (with water jacket, 2) with an oxygen electrode inserted (3) and magnetic stirrer coupled (4). The electrode determines O2 concentration in aqueous solutions over a period of time. (b) Oxygen electrode inset (from ref. 3) The electrode itself is located inside the chamber, in horizontal position (or depending the general apparatus design). The platinum cathode (5) is located surrounding a rod-like center anode (6) made by silver (reference Ag/AgCl electrode). These two electrodes are connected with each other by a thin layer of electrolyte (50% saturated KCl-solution, 7). Directly placed on top of the rod, there is an oxygen-permeable Teflon membrane (8), that is tight fitted by a rubber ring (9). The O2 recordings can be done in open or closed chamber mode. In the last case, the reciprocal air-solution O2 diffusion is avoided which allows for better determination of the respiratory rates. Experimental additions of solutions, substrates, mitochondria, and substrates/inhibitors are done through the top of the chamber (10) (open mode) or through the small hole (11) inside a stopper, respectively.

how intact or how coupled mitochondria are. The initial step to obtain RCR is to independently calculate state 3 and state 4 respiration rates. State 3 respiration is triggered in a suspension of isolated mitochondria by the addition of ADP. The increase in respiration denotes the use of the proton-motive force for the synthesis of ATP, being restored by the augmented proton pumping activity of the respiratory chain. When all ADP is phosphorylated into ATP by the action of the ATP-synthase, the respiration returns to or close to the initial pre-ADP addition values. The new respiratory state is now termed state 4. Some authors describe other respiratory states including state 2, which is the mitochondrial respiration in the presence of substrate only (pre-ADP addition), while others even establish a respiratory state 1, when mitochondria are consuming oxygen by oxidizing internal substrates only. Regardless of the respiratory state being measured, units are usually expressed as nmol O2/min/mg protein or natom O/min/mg protein.

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There are several important values to take into account when calculating the rates of oxygen consumption. By the units involved, it is clear that the exact time in which an x amount of oxygen is consumed is critical. Also, the amount of mitochondrial protein used in the experiment must be well known. There are several precise protein quantification methods including Bradford and Biuret methods, among others (20, 21). For cells, the oxygen consumption may be rationalized for the number of cells considered in oxygen reaction chamber, and data present in function of nmol O2/min/cells number (cell number may be quantified, depending from the biological samples) or when considering the total protein, we can use the method described by Lowry et al. (13). Some authors may actually normalize oxygen consumption rates to the activity of citrate synthase, to normalize for mitochondrial content (22, 23) (see Note 3). If oxygen consumption is being measured in a flatbed recorder, one important parameter that should be taken into account is the scale of the paper, which will determine which distance in the paper will correspond to 100% oxygen saturation in water or in saline (several tables can be consulted for the values at the desired temperature (see Notes 4 and 5)). Also, paper recording velocity, which is adjustable, must be recorded even before the assays start. In this regard, it is advised to reach a compromise between a slow paper velocity, which would allow for a better measurement of the rates or a faster paper velocity, which turns the measurement of the faster rates harder but allows to spare the researcher the waste of long stretches of paper. After measuring state 3 and state 4 respiration rates, the RCR is easily calculated by performing the rate between state 3 and state 4. The final value should not present any units. There is a wide variety of tables showing values that are consistent with a good mitochondrial preparation (for example, see ref. 4). As an example, we routinely measure the RCR of isolated mitochondria in our laboratory just after the isolation or when investigating the effects of different xenobiotics. We usually have RCRs in the order of 6–8 for complex I substrates and 2–4 for complex II substrates (we have slightly lower values for heart vs. liver, most likely because of higher intrinsic ATPase activity in our heart mitochondrial fractions, which lead to a higher state 4 respiration). When using ascorbate plus N,N,N¢,N¢-tetramethyl-p-phenylenediamine (TMPD) to directly feed complex IV via cytochrome c, the RCR value is around 1.5–2, which occurs due to the very fast respiration during state 4 respiration, since only one proton pump is working (complex IV or cytochrome c oxidase). Damaged mitochondria usually have low RCRs, being 1, the theoretical minimum value. One test compound will decrease the RCR as it leads to increased state 4 respiration, decreased state 3 respiration or both.

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The ADP/O has a completely different meaning. This parameter measures the efficiency of the mitochondrial phosphorylative system; in other words, the ADP/O index, which has no units, will give us a measure of how much oxygen the respiratory chain reduces to water per ADP phosphorylated. The index is calculated as the number of nmol of ADP added to the system divided by the number of natoms oxygen consumed during state 3 respiration. And this last point is critical, since a common mistake is to calculate the ADP/O as the amount of ADP added in nmol divided by the respiration rate during state 3. After measuring the absolute oxygen consumption and converting it into natoms, the calculation is now easy to do. One common strategy to calculate the amount of oxygen during state 3 respiration is to draw lines (as shown in Fig. 3) on top of the respiration curves during states 2, 3, and 4 and use the intersection points between state 2/3 and state 3/4 as a measure to calculate the absolute distance in the paper, parallel to the axis corresponding to oxygen concentration in the media. There is a large controversy on the theoretical values for the ADP/O value (24), since the ADP/O is dependent on the coupling ratios of proton transport. New discoveries on the structure and activity of the ATP-synthase justify a new assessment of the established values, also because several other mechanisms account for the different ADP/O values found in the literature (24, 25). Under our conditions, we routinely have values around 2.4–3.1 for

Fig. 3. Representative oxygen consumption recording (in a flatbed recorder) from mitochondrial respiration assays, using Clark-type electrode. Dashed lines (a, b, and c) were drawn (for plot 1, as an example) for ADP/O and RCR calculations (see text body). (a) Respiratory plots in the presence of glutamate/malate; (b) respiratory plots in the presence of succinate.

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complex I substrates and 1.4–1.9 for complex II substrates. Also common during bench work is the appearance of ADP/O values much higher or lower than expected. The researcher must be aware that an incorrectly measured ADP stock solution will lead to an incorrect determination of ADP/O values (see Note 6). Finally, one common source of error is when the oxygen electrode used is not responding fast enough to the alterations in the media oxygen content. When this happens, the transition between each state is slow and does not have an abrupt profile, which will lead to the underestimation of the ADP/O value. A good and fast-responding oxygen electrode is thus essential for correct estimate of the index (see Note 7). Also, membranes need to have good O2 permeability properties (see Note 8). In oxygen consumption experiments, we can also consider other mitochondrial respiration states (7), such as “state FCCP” (uncoupled respiration, maximal activity of the respiratory chain) or “state oligomycin”, which is usually used to determine oxygen consumption with inhibited ATP synthase. 3.6. Oxygen Consumption Assays: Permeabilized Animal Cells

Oxygen consumption assessment implies the choice of the bestsuited reaction chamber, accordingly with the biological preparation volumes. For all types of presented cells we need to consider: 1. Check if temperature in reaction/O2 consumption chamber is 37°C. 2. Calibrate O2 scale using distilled water. 3. Introduce the desired volume of respiration buffer, and just place biological preparations after reaching a steady baseline on the chart. 4. In electropermeabilized, digitonin-permeabilized or saponinpermeabilized cells, a schematic sequence of additions (using for example microsyringes) such as 10 mM pyruvate + 5 mM malate, 10 mM glutamate + 5 mM, 2 mM ascorbate + TMPD and 2.5  mM FCCP (see Note 1) and the addition of ADP (125–250 nM) after a substrate can be used to check OXPHOS function.

3.7. Oxygen Consumption Assays: Isolated Mitochondria

1. Define appropriate temperature, usually between 25 and 30°C. 2. Calibrate appropriate O2 scale using distilled water (for example, a 1.5× scale can be appropriate in most cases to get a better signal resolution for isolated mitochondria). 3. Introduce the desired volume of respiration buffer, and just place biological preparations after reaching a steady baseline on the chart. 4. Add mitochondrial suspension with the desired protein concentration.

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Sequential additions of mitochondrial substrates and inhibitors can allow characterizing different segments of the respiratory chain in one experiment only. We can use a schematic sequence of additions such as 10 mM pyruvate + 5  mM malate followed by ADP (125–250  nmol) and later rotenone (2 mM), followed by 5 mM succinate, ADP (same as before), with antimycin (2  mM) later on. If still far from total chamber oxygen consumption, a subsequent addition can be performed with 2  mM ascorbate + TMPD, plus ADP. Another possibility is the addition of 1 mM FCCP (see Note 1, Table 1) to uncouple respiration. The number of additions depends on the oxygen availability in the chamber and enough time should be elapsed between additions to warranty correct measurement of oxygen consumption rates.

Table 1 Most-used substrates and inhibitors during oxygen consumption assays Name

Classification

Place of action

Antimycin A

Inhibitor

Complex III

Ascorbate

Substrate

Complex IV (by donating electrons to cytochrome c)

Atractyloside

Inhibitor

Adenine nucleotide translocator (no ADP/ATP exchange occurs)

FCCP

Protonophore and uncoupler of mitochondrial OXPHOS

Mitochondrial membrane (although proton shuttling activity can be facilitated by some mitochondrial proteins)

Fumarate

Substrate

Intermediate in Krebs cycle

Glutamate

Substrate

Complex I (converted into NADH in the matrix)

KCN

Inhibitor

Complex IV

Malate

Substrate

Complex I (converted into NADH in the matrix)

Malonate

Inhibitor

Inhibition of succinate oxidation

Myxothiazol

Inhibitor

Complex III

Oligomycin

Inhibitor

Complex V

Pyruvate

Substrate

Feeds Krebs cycle (via Acetyl-CoA)

Rotenone

Inhibitor

Complex I

Succinate

Substrate

Complex II

TMPD

Reducing co-substrate

By-passing electrons to Complex IV (via cytochrome c), as an artificial electron mediator

FCCP carbonylcyanide p-trifluoromethoxyphenylhydrazone, KCN potassium cyanide, TMPD N,N,N ¢,N ¢-tetramethylp-phenylenediamine

2  Evaluation of Respiration with Clark Type Electrode…

21

4. Notes 1. During oxygen consumption, assays to follow mitochondrial or cell respiration must be considered particular substrates and/or inhibitors for individual respiratory complexes assessment (see Table 1). 2. Electric circuitry that controls oxygen electrode. One tension division circuit (inside galvanometer) generates an electrode potential difference of 0.71  V (between anode and cathode) which is proportional to the activity of water-dissolved O2 solution (16). 3. By measuring the latent citrate synthase activity (calculating the difference between total citrate synthase activity and free citrate synthase activity) or citrate synthase ratio (latent citrate synthase activity/free citrate synthase activity), it is possible to infer not only the structural integrity of the mitochondrial preparation but also to correlate oxygen consumption with the quantity of the functional mitochondria or cells with functional mitochondria, instead of the total protein. Rationing oxygen consumption by citrate synthase content is time-consuming and thus rarely used. 4. Oxygen solubility differs according to the type of media we use. We can find many tabled values for distilled water and different saline buffers (26, 27) but our experience tells us that a good approach is to use standard oxygen-saturated deionised water for calibration. 5. The most correct step to calibrate the O2 scale is by comparing the signal of air-saturated water with the zero level created by adding dithionite to the water in the reaction chamber (this starts a reaction that quickly consumes the dissolved oxygen) or flushing the cell with nitrogen (17). But when comparing oxygen consumption from specific conditions to a control, knowing or not, the precise values of dissolved O2 will not have a meaningful impact on data analysis. 6. Spectrophotometric determination of ADP stock solutions should always be conducted before start experiments with a new prepared batch. ADP concentration can be determined by measuring the absorbance of the batch at 259  nm (molar extinction coefficient = 15.4 × l03/M cm) (28). 7. To maintain the electrode in good conditions, it should be cleaned after use and before prolonged storage. Never allow the electrode to dry without the electrolyte in place, as crystallization of the electrolyte may occur and cause the platinum/ epoxy resin seal to be breached and allow the deposition of electrolyte salt crystals around the cathode, leading to necessary

22

A.M. Silva and P.J. Oliveira

electrode replacement. The silver electrode (anode) must be cleaned to remove the excessive deposits of black oxides, which could cause deterioration. For that, we use a small cotton bud moistened with distilled water. Some commercial products can be used with the purpose of cleaning silver electrodes, but avoid any harsh abrasive substance. Platinum cathode cleaning involves the usage of a non-corrosive polishing agent (as a soft white tooth paste, with small silica spheres in the composition) on top of a cotton bud moistened with few drops of distilled water. Circular movements in the polish platinum electrode surface can be performed to obtain a final shinny effect. Make sure that all electrical connections are preserved and completely dry (29). 8. Several different membrane materials can be used for the oxygen electrode including Teflon, polyethylene, and silicon rubber, among others. Thin hydrophilic polymers are usually not mechanically strong which may cause problems. It is possible to obtain some degree of selectivity by choosing the material of this membrane according to the conditions of application. The diffusion through such a structure is more complicated. For radial geometry, the steady-state current, I, is given by the mathematical expression:

4π F0 Ds Ss P(r1 ) I= Ds (r1 − r0 ) / Dm r1 + Sm r0 r1



D corresponds to the diffusion coefficient (in solution and in the membrane), r0 is the outer radius of the membrane, and r1 is the radius of the electrode. S represents oxygen solubility in solution and in the membrane (Ss and Sm, respectively). Another important parameter is the oxygen partial pressure on the membrane pressure, P(r1) (6). Literature reports a wide range of membranes for Clark-type oxygen sensors. Jobst et  al. (30) considered a membrane based on a thin-layer polymer, to prepare a planar oxygen sensor with a three-electrode configuration (platinum working and counter electrodes and Ag/AgCl reference electrode) that allows the regeneration of oxygen consume in the cathode. The electrolyte is based in a hydrogel layer, preventing the typical buffer degradation and self oxygen consuming behavior. But for mitochondrial preparations, the system is not the best suited to be coupled to the common oxygen recording chamber, and membrane preparation requires a complex experimental setup.

2  Evaluation of Respiration with Clark Type Electrode…

23

Acknowledgments Work in the author’s laboratory is supported by the Portuguese Foundation for Science and Technology (research grants PTDC/ QUI-QUI/101409/2008 and PTDC/AGR-ALI/108326/2008). References 1. Du G, Mouithys-Mickalad A, Sluse FE (1998) Generation of superoxide anion by mitochondria and impairment of their functions during anoxia and reoxigenation in  vitro. Free Rad Biol Med 25(9):1066–1074 2. Stryer L (1999) Biochemistry, 4th edn. W. H. Freeman, New York 3. Severinghaus JW, And Astrup PB (1986) History of blood gas analysis. IV. Leland Clark’s oxygen electrode. J Clin Monit 2(2):125–139 4. Pereira GC, Branco AF, Matos JA, Pereira SL, Parke D, Perkins EL, Serafim TL, Sardão VA, Santos MS, Moreno AJ, Holy J, Oliveira PJ (2007) Mitochondrially targeted effects of Berberine [natural yellow 18,5,6-dihydro-9,10dimethoxybenzo(g)-1,3-benzodioxolo(5,6-a) quinolizinium] on K1735-M2 mouse melanoma cells: comparison with direct effects on isolated mitochondrial fractions. J Pharmacol Exp Ther 323(2):636–649 5. Estabrook RW (1967) Mitochondrial respiratory control and the polarographic measurement of ADP:O ratios. Methods Enzimol 10:41–47 6. Janata J (2009) Principles of chemical sensors, 2nd edn. Springer, New York 7. Respiratory states and coupling control ratios. In: Mitochondrial pathways and respiratory control, 2nd edn. Oroboros MiPnet Publications, Innbruck 8. Wenchich L, Drahota Z, Honzík T, Hansíková, Tesařová M, Zeman J, Houštĕk J (2003) Polarographic evaluation of mitochondrial enzymes activity in isolated mitochondria and in permeabilized human muscle cells with inherited mitochondrial defects. Physiol Res 52:781–788 9. Grinstein S, Furuya W, Lu DJ, Mills GB (1990) Vanadate stimulates oxygen consumption and tyrosine phosphorilation in electropermeabilized human neutrophils. J Biol Chem 265:318–327 10. Conget I, Barrientos A, Manzanares JM, Casademont J, Viñas O, Barceló J, Nunes V, Gomis R, Cardellach F (1997) Respiratory chain activity and mitochondrial DNA content

of nonpurified and purified pancreatic islet cells. Metabolism 46(9):984–987 11. Artuch R, Colomé C, Playán A, Alcaine MJ, Briones P, Montoya J, Vilaseca MA, Pineda M (2000) Oxygen consumption measurement in lymphocytes for the diagnosis of pediatric patients with oxidative phosphorylation diseases. Clin Biochem 33(6):481–485 12. Rolo AP, Palmeira CM, Cortopassi GA (2009) Biosensor plates detect mitochondrial physiological regulators and mutations in  vivo. Anal Biochem 385(1):176–178 13. Lowry OH, Rosenbrough NJ, Far AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193(1):265–275 14. Moreira PI, Santos MS, Moreno AM, Seica R, Oliveira CR (2003) Increased vulnerability of brain mitochondria in diabetic (Goto-Kakizaki) rats with aging and amyloid-b exposure. Diabetes 52:1440–1456 15. Alves MJ, Oliveira PJ, Carvalho RA (2009) Mitochondrial preservation in celsior versus histidine buffer solution during cardiac ischemia and reperfusion. Cardiovasc Toxicol 9(4): 185–193 16. Moreno AJM (1992) Estudo do efeito de Insecticidas de usos comum na bioenergética mitocondrial. Universidade de Coimbra, Tese de Doutoramento 17. Renger G, Hanssum B (2009) Oxygen detection in biological systems. Photosynth Res 102:487–498 18. Diepart C, Verrax J, Calderon PB, Feron O, Jordan BF (2010) Comparison of methods for measuring oxygen consumption in tumor cells in vitro. Anal Biochem 396:250–256 19. Kopustinskas A, Adaskevicius R, Krusinskas A, Kopustinskiene DM, Liobikas J, Toleikis A (2006) A user-friendly PC-based data acquisition and analysis system for respirometric investigations. Comp Method Prog Biomed 82:231–237 20. Gornall AG, Bardawill CS, David MM (1949) Determination of serum proteins by means of the biuret reaction. J Biol Chem 177: 751–766

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2 1. Bradford MM (1976) A rapid and sensitive for the quantitation of microgram quantities of protein utilizing the principle of protein– dye binding. Anal Biochem 72: 248–254 22. Bugger H, Chemnitius J, Doenst T (2006) Differential changes in respiratory capacity and ischemia tolerance of isolated mitochondria from atrophied and hypertrophied hearts. Metab Clin Exp 55:1097–1106 23. Figueiredo PA, Ferreira RM, Appell HJ, Duarte JA (2008) Age-induced morphological, biochemical, and functional alterations in isolated mitochondria from murine skeletal muscle. J Gerontol A Biol Sci Med Sci 63: 350–359 24. Hinkle PC (2005) P/O ratios of mitochondrial oxidative phosphorylation. Biochim Biophys Acta 1706(1–2):1–11 25. Lee CP, Gu Q, Xiong Y, Mitchell RA, Ernster L (1996) P/O ratios reassessed: mitochondrial P/O ratios consistently exceed 1.5 with succinate

and 2.5 with NAD-linked substrates. FASEB J 10(2):345–350 26. Atkins PW (1998) Physical chemistry, 6th edn. Oxford University Press, New York 27. Rasmussen HN, Rasmussen UF (2003) Oxygen solubilities of media used in electrochemical respiration measurements. Anal Biochem 319:105–113 28. Young M (1967) On the interaction of adenosine diphosphate with myosin and its enzymatically active fragments. J Biol Chem 242(11):2790 29. Hansatech Instruments (2006) Operations manual – setup, installation and maintenance. Electrode preparation and maintenance. Version 2.2. Hansatech Instruments Ltd, Norfolk 30. Jobst G, Urban G, Jachimowicz A, Kohl F, Tilado O, Lettenbichler I, Nauer G (1993) Thin-film Clark-type oxygen sensor based on novel polymer membrane systems for in  vivo and biosensor applications. Biosens Bioelectron 8(3–4):123–128

Chapter 3 High-Resolution Respirometry: OXPHOS Protocols for Human Cells and Permeabilized Fibers from Small Biopsies of Human Muscle Dominik Pesta and Erich Gnaiger Abstract Protocols for high-resolution respirometry (HRR) of intact cells, permeabilized cells, and permeabilized muscle fibers offer sensitive diagnostic tests of integrated mitochondrial function using standard cell culture techniques and small needle biopsies of muscle. Multiple substrate–uncoupler–inhibitor titration (SUIT) protocols for analysis of oxidative phosphorylation improve our understanding of mitochondrial respiratory control and the pathophysiology of mitochondrial diseases. Respiratory states are defined in functional terms to account for the network of metabolic interactions in complex SUIT protocols with stepwise modulation of coupling and substrate control. A regulated degree of intrinsic uncoupling is a hallmark of oxidative phosphorylation, whereas pathological and toxicological dyscoupling is evaluated as a mitochondrial defect. The noncoupled state of maximum respiration is experimentally induced by titration of established uncouplers (FCCP, DNP) to collapse the proton gradient across the mitochondrial inner membrane and measure the capacity of the electron transfer system (ETS, open-circuit operation of respiration). Intrinsic uncoupling and dyscoupling are evaluated as the flux control ratio between nonphosphorylating LEAK respiration (electron flow coupled to proton pumping to compensate for proton leaks) and ETS capacity. If OXPHOS capacity (maximally ADP-stimulated oxygen flux) is less than ETS capacity, the phosphorylation system contributes to flux control. Physiological Complex I + II substrate combinations are required to reconstitute TCA cycle function. This supports maximum ETS and OXPHOS capacities, due to the additive effect of multiple electron supply pathways converging at the Q-junction. Substrate control with electron entry separately through Complex I (pyruvate + malate or glutamate + malate) or Complex II (succinate + rotenone) restricts ETS capacity and artificially enhances flux control upstream of the Q-cycle, providing diagnostic information on specific branches of the ETS. Oxygen levels are maintained above air saturation in protocols with permeabilized muscle fibers to avoid experimental oxygen limitation of respiration. Standardized two-point calibration of the polarographic oxygen sensor (static sensor calibration), calibration of the sensor response time (dynamic sensor calibration), and evaluation of instrumental background oxygen flux (systemic flux compensation) provide the unique experimental basis for high accuracy of quantitative results and quality control in HRR. Key words: Substrate–uncoupler–inhibitor titration, Human vastus lateralis, Needle biopsy, HEK, HPMC, HUVEC, Fibroblasts, Routine respiration, Oxidative phosphorylation, Q-junction, Pyruvate, Glutamate, Malate, Succinate, Leak, Coupling control, Uncoupling, Oxygraph, Oxygen flux, Residual oxygen consumption, Instrumental background

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_3, © Springer Science+Business Media, LLC 2012

25

26

D. Pesta and E. Gnaiger

Abbreviations CCP E FCR HRR L mt O2k P POS R ROX SUIT Ww

Coupling control protocol Electron transfer system capacity Flux control ratio High-resolution respirometry LEAK respiration Mitochondrial Oxygraph-2k OXPHOS capacity Polarographic oxygen sensor ROUTINE respiration Residual oxygen consumption Substrate–uncoupler–inhibitor titration Wet weight

1. Introduction Mitochondrial respiration is a key element of cell physiology. Cell respiration channels metabolic fuels into the bioenergetic machinery of oxidative phosphorylation, regulating and being regulated by molecular redox states, ion gradients, mitochondrial membrane potential, the phosphorylation state of the ATP system, and heat dissipation in response to intrinsic and extrinsic energy demands. Complementary to anaerobic energy conversion, mitochondrial respiration is the aerobic flux of life. It integrates and transmits a wide range of physiological and pathological signals within the dynamic communication network of the cell. This provides the background for an interdisciplinary interest in the study of mitochondrial respiratory function in biology and medicine (see Note 1). Respirometry reflects the function of mitochondria as structurally intact organelles. It provides a dynamic measurement of metabolic flux (rates), in contrast to static determination (states) of molecular components, such as metabolite and enzyme levels, redox states and membrane potential, concentrations of signaling molecules, or RNA and DNA levels. Mitochondrial respiratory function, therefore, cannot be measured on frozen tissue samples, but usually requires minimum storage times of biological samples and delicate handling procedures to preserve structure and function. Moreover, mitochondrial respiration yields an integrative measure of the dynamics of complex coupled metabolic pathways, in contrast to monitoring activities of isolated enzymes. Measurement of respiratory flux in different metabolic states is required for evaluation of the effect on oxidative phosphorylation of changes in metabolite levels, membrane permeability, or activity of individual enzymes. Small imbalances in metabolic flux can result in large cumulative

3

High-Resolution Respirometry: OXPHOS Protocols…

27

changes of state of a metabolic system. Vice versa, even a large defect of individual enzymes may result in minor changes of flux, due to threshold effects. Hence high-quantitative resolution of respirometry is required for diagnostic applications, particularly when amounts of cells or tissue are limiting. Understanding mitochondrial respiratory control, in turn, requires experimental modulation of metabolite levels, electrochemical potentials, and enzyme activities. HRR has been developed to meet these demands and to provide the instrumental basis for modular extension with additional electrochemical and optical sensors for investigations of mitochondrial respiratory physiology (1–3). This chapter describes applications of HRR with intact and permeabilized cells, and permeabilized muscle fibers for functional mitochondrial diagnosis. Protocols are presented for cell membrane permeabilization, biopsy sampling, short-term storage, fiber preparation, and respirometric titration regimes. The protocols provide diagnostic tests for evaluation of membrane integrity (coupling of oxidative phosphorylation; cytochrome c release), respiratory inhibition resulting from defects in the phosphorylation system or ETS, including respiratory complexes and activities of dehydrogenases and metabolite transporters across the inner mitochondrial membrane (4). Quality control in HRR includes traceability of basic sensor calibration, systemic flux compensation, kinetic evaluation of ADP saturation, and oxygen dependence of respiration with permeabilized fibers (2, 5), and integrates the general features of the OROBOROS Oxygraph-2k.

2. Materials 2.1. Materials for Preparation of Permeabilized Muscle Fibers

1. Nonmagnetic forceps are used for fiber preparation, one pair with sharp straight tips; one pair with sharp rounded tips; two pairs with very sharp angular tips.

2.2. Media and Chemicals

1. Mitochondrial respiration medium (MiR05), 110 mM sucrose, 60 mM K+-lactobionate, 0.5 mM EGTA, 3 mM MgCl2, 20 mM taurine, 10 mM KH2PO4, 20 mM HEPES adjusted to pH 7.1 with KOH at 37 °C; and 1 g/l BSA essentially fatty acid free (6). MiR06 is MiR05 plus 280 U/ml catalase (7).

2. Microbalance with five digits; 0.01 mg (Mettler-Toledo XS105DU or XS205DU) for the measurement of wet weight of fibers.

2. Relaxing and biopsy preservation solution for muscle fibers (BIOPS), 50 mM K+-MES, 20 mM taurine, 0.5 mM dithiothreitol, 6.56 mM MgCl2, 5.77 mM ATP, 15 mM phosphocreatine, 20 mM imidazole, pH 7.1, adjusted with 5 N KOH at 0 °C,

28

D. Pesta and E. Gnaiger

10 mM Ca–EGTA buffer (2.77 mM CaK2EGTA + 7.23 mM K2EGTA; 0.1 mM free calcium; ref. 8). ATP is hydrolyzed at least partially during fiber storage, thus generating millimolar levels of inorganic phosphate. It has not been tested if addition of 3 mM phosphate (9) exerts any effect on preservation quality. BIOPS can be stored at −20 °C. 3. Selected substrates, uncouplers, and inhibitors are listed in Table 1, with corresponding Hamilton syringes used for manual titrations into a 2-ml Oxygraph chamber. The sources of chemicals change according to availability and evaluation of quality and price. Information is summarized and can be updated on http://www.bioblast.at. 2.3. The Oxygraph-2k for HRR

New methodological standards have been set by HRR with the OROBOROS Oxygraph-2k (O2k; Fig. 1; OROBOROS INSTRUMENTS, Austria; http://www.oroboros.at; refs. 1–3, 10). The principle of closed-chamber HRR is based on monitoring oxygen concentration, cO2 , in the incubation medium over time, and plotting oxygen consumption by the biological sample ( J O2 ) and cO2 continuously while performing the various titrations in a respirometric protocol (Figs. 2 and 5–8). Without compromise on HRR features, the O2k provides robustness and reliability of routine instrumental performance. To increase throughput particularly in research with cell cultures and biopsy samples, the user-friendly integrated concept with full software support (DatLab) makes it possible to apply several instruments in parallel, each O2k with two independent chambers (Fig. 1). Chambers, sensors, and the electronics are shielded by a copper block and stainless steel housing. Angular insertion of the oxygen sensor into the cylindrical glass chamber places the polarographic oxygen sensor into an optimum position for stirring (Fig. 1, inset). Integrated electronic control includes Peltier temperature regulation (2–45 °C, stability ±0.001 °C), stirrer control, an electronic barometric pressure transducer for air calibration, and the optional automatic titration-injection micropump TIP2k (Fig. 1).

3. Methods 3.1. Respirometry with Intact Cells: Coupling Control Protocol

Aerobic and anaerobic metabolism is physiologically controlled in the ROUTINE state of cell respiration. Different coupling control states (4) are induced by the application of membrane-permeable inhibitors and uncouplers in a coupling control protocol (CCP; Table 2). 0.3–1.0 million fibroblasts or endothelial cells per experiment are sufficient using the OROBOROS Oxygraph-2k at 37 °C (1–3). Cell densities are adjusted to obtain maximum fluxes in the

M

G

S

Oct

As

Malate

Glutamate

Succinate

Octanoylcarnitine

Ascorbate

c

D

T

F F

Rot Mna Ama

Cyt.

ADP

ATP

Uncoupler FCCPb FCCP

Inhibitors Rotenone Malonic acid Antimycin A

TMPD

Tm

P

Pyruvate

a

Abbr.

Substrates

CI CII CIII

DmH+ DmH+

CV, ANT

CV, ANT

CIV

CIV

CIV

ETF

CII

CI

CI

CI

Site of action

1.0 mM (EtOH) 2 M (H2O) 5 mM (EtOH)

1 mM (EtOH) 0.1 mM (EtOH)

0.5 M (H2O)

−20 Fresh −20

−20 −20

−80

−80

−20

4 mM (H2O) 0.5 M (H2O)

−20

−20

−20

−20

−20

−20

Fresh

Storage (°C)

0.2 M (H2O)

0.8 M (H2O)

0.1 M (H2O)

1 M (H2O)

2 M (H2O)

0.8 M (H2O)

2 M (H2O)

Concentration in syringe (solvent)

0.5 mM 5.0 mM 2.5 mM

0.5 mM steps 0.05 mM steps

1–5 mM

1–5 mM

10 mM

0.5 mM

2 mM

0.2 mM

10 mM

10 mM

2 mM

5 mM

Final concentration in 2-ml O2k-chamber

1 5

1 ml steps 1 ml steps

4–20

4–20

5

5

5

4

20

10

5

5

Titration (ml) into 2 ml

(continued)

10 25 10

10 10

10

10

25

25

25

10

50

25

25

25

Syringe (ml)

Table 1 Selected substrates, uncouplers, and inhibitors used in SUIT protocols with isolated mitochondria or permeabilized fibers, with abbreviations (Abbr.), and site of action (electron entry, substrate entry, or inhibition of ATP synthase, CV; adenine nucleotide translocase, ANT) (37)

3 High-Resolution Respirometry: OXPHOS Protocols… 29

Myx Azd Kcn Omy Atr

Myxothiazol Sodium azide KCN Oligomycin Atractyloside

CIII CIV CIV CV ANT

Site of action 1 mM (EtOH) 4 M (H2O) 1 M (H2O) 4 mg/ml (EtOH) 50 mM (H2O)

Concentration in syringe (solvent)

b

N,N,N,N ¢,N ¢-tetramethyl-p-phenylenediamine dihydrochloride Carbonyl cyanide p-trifluoromethoxyphenylhydrazone

a

Abbr.

Substrates

Table 1 (continued)

−20 −20 Fresh −20 −20

Storage (°C) 0.5 mM ³100 mM 1.0 mM 2 mg/ml 0.75 mM

Final concentration in 2-ml O2k-chamber

1 ³50 2 1 30

Titration (ml) into 2 ml

10 50 10 10 50

Syringe (ml)

30 D. Pesta and E. Gnaiger

Peltier temperature control, ±0.001°C O2k Stainless steel housing

Titration-Injection microPump TIP2k Syringe A

O2 Sensor POS A

Needle A

Stopper A ISS

Window Chamber A O2 Sensor POS A

A

B

Stopper

Insulated copper block POS

Pre-amplifyer POS B

Stirrer control light B

PVDF

Ion sensitive electrode plug B

stirrer Glass chamber

Barometric pressure transducer

Fig. 1. Oxygraph-2k for high-resolution respirometry (O2k Series C) with TIP2k and integrated suction system (ISS). Two glass chambers (A and B) are housed in an insulated copper block with electronic Peltier temperature control. Polarographic oxygen sensors (POS) are sealed by a butyl rubber gasket against the angular plane on the glass chambers. The PVDF (or PEEK) stirrers are powered by electrically pulsed magnets inserted in the copper block. Stoppers contain a capillary for extrusion of gas bubbles and insertion of a needle for manual or automatic titrations with the TIP2k. Additional capillaries through the stopper (PVDF) are drilled for insertion of various electrodes, the signals of which are simultaneously recorded by the DatLab software. Copyright ©2010 by OROBOROS INSTRUMENTS. Reproduced with permission; http://www.oroboros.at.

LEAK ETS

ROX

O2 concentration, c O2 [µM]

200 S+D

Omy

FCCP

Rot

Ama

ES

150

150

c O2 100

100 I O2

RS

R

50

50 LS

ROXS

ROX 0

0 0

10

20 30 Time [min]

40

50

Respiration, IO2 [pmol·s−1·10−6 cells]

ROUTINE 200

Fig. 2. Coupling control protocol and respirometric cell viability test, showing continuous traces of respiration of human peritoneal mesothelial cells [ I O (pmol/s per 106 cells)], calculated as the negative time derivative of oxygen concentration, 2 cO2 . 0.5 × 106 cells/ml in MiR05 (80% viability; 2-ml chamber, 37 °C). Endogenous ROUTINE respiration without substrates (R), addition of 10 mM succinate and 5 mM ADP (S + D), 1 mg/ml oligomycin (Omy; LEAK state, L), stepwise 1 mM titration of FCCP (ETS capacity, state E), and inhibition by 0.5 mM rotenone (Rot) and 2.5 mM antimycin A (Ama) for the final measurement of residual oxygen consumption (ROX). Oxygen flow is not shown immediately after titrations, when oxygen concentration and the calculation of slopes are disturbed particularly when ethanol is the solvent. Modified after ref. 19.

54 ± 11

CEM – Controld

Fibroblasts – Senescent

6

285 ± 72

138 ± 22

Fibroblasts – Young arrestg

g

111 ± 24

142 ± 47

Fibroblasts – Youngg

HPMC + IL-1b

f

HPMC Control

181 ± 58

114 ± 18

HUVECe

f

85 ± 13

CEM – S-phase, apopt.d

CEM – G1-phase, apopt.

0.42 ± 0.05

0.23 ± 0.01

0.34 ± 0.03

0.41 ± 0.09

0.40 ± 0.09

0.26 ± 0.02

0.38 ± 0.03

0.41 ± 0.03

0.40 ± 0.03

0.39 ± 0.02

0.31 ± 0.03

ROUTINE, R/E

0.21 ± 0.04

0.05 ± 0.01

0.14 ± 0.02

0.08 ± 0.02

0.09 ± 0.01

0.13 ± 0.02

n.d.

n.d.

n.d.

0.10 ± 0.02

0.09 ± 0.00

LEAK, L/E

0.21 ± 0.05

0.18 ± 0.02

0.20 ± 0.02

0.32 ± 0.08

0.31 ± 0.08

0.13 ± 0.00

n.d.

n.d.

n.d.

0.29 ± 0.02

0.23 ± 0.02

netROUTINE, (R − L)/E

0.07 ± 0.03

0.05 ± 0.00

0.07 ± 0.03

0.02 ± 0.01

0.005 ± 0.01

0.05 ± 0.04

0.01 ± 0.01

0.03 ± 0.03

0.02 ± 0.03

0.03 ± 0.01

0.01 ± 0.00

Residual, ROX/E ¢

(15)

(15)

(15)

(19)

(19)

(16)

(12)

(12)

(12)

(3)

(17)

References

Capacity of the ETS (pmol/s per 10 cells) is the reference for normalization of FCR, E = E ¢ − ROX, where E¢ i s the apparent (uncorrected) electron transfer capacity. Similarly, R = R¢ − ROX and L = L¢ − ROX. To calculate total ROUTINE respiration (R¢ (pmol/s per 106 cells)) R ¢ = ((R /E ) + (ROX / E ¢) / (1 - ROX / E ¢)) ´ E b Transformed human embryonic kidney cells (10 × 106/ml); N = 3–8 independent cell cultures in culture medium DMEM c Mouse parental hematopoietic cells (1.1 × 106/ml); n = 6, replicate O2k measurements of a single suspension culture in culture medium RPMI d Human leukemia cells (1.0 × 106/ml to 1.2 × 106/ml); controls (N = 27), and 30% apoptotic cultures preincubated with dexamethasone, arrested in the G1-phase (N = 9) or with gemcitabine, arrested in the S-phase (N = 12) in culture medium RPMI; n.d., not determined e Human umbilical vein endothelial cells (0.9 × 106/ml); N = 3; in culture medium EGM f Human peritoneal mesothelial cells (0.6 × 106/ml); N = 5; cultured from five donors, incubated for 48 h without (controls) or with recombinant IL-1b. ROUTINE respiration in MiR05 with succinate and ADP (Fig. 2) g Human foreskin fibroblasts; young (n = 12; 1.0 × 106/ml), young-cell cycle arrest (n = 5; 1.1 × 106/ml), and senescent (n = 12; 0.2 × 106/ml), in culture medium DMEM

a

81 ± 11

32D (mouse)c

31 ± 6

14 ± 2

HEK 293b

d

ETS, E (pmol/s/106)

Cell type

Table 2 Electron transfer system (ETS) capacity and flux control ratios (FCRs)a in the coupling control protocol with intact human (and 32D mouse) cells (37 °C; mean ± SD)

32 D. Pesta and E. Gnaiger

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range of 30–300 pmol/s ml. It takes 50–90 min for the evaluation of mitochondrial coupling states (Fig. 2). Respiratory flow, I O2 , of intact cells is expressed per million viable cells (Table 2). Cell viability should be >0.95 in the control group of various cell types (HUVEC, fibroblasts). Mass-specific oxygen flux, J O2 , is expressed per mg dry weight (11) or cell protein (12). Mitochondrial (mt) marker-specific respiration is obtained by normalization of flux relative to a mt-marker, such as citrate synthase (CS) or cytochrome c oxidase activity (12). Internal normalization yields flux control ratios relative to flux in a common reference state (Table 2). 3.1.1. ROUTINE Respiration

Cellular ROUTINE respiration (R) and growth is supported by exogenous substrates in culture media (3, 11–17). In media without energy substrates, respiration is based on endogenous substrates. Physiological energy demand, energy turnover, and the degree of coupling (intrinsic uncoupling and pathological dyscoupling) control the levels of respiration and phosphorylation in the physiological coupling state R of intact cells. The maximum capacity of oxidative phosphorylation (OXPHOS; analogous to State 3; ref. 18) cannot be studied by external addition of ADP to intact cells, since the plasma membrane is impermeable to ADP and many mitochondrial substrates (11–14, 19). However, evaluation of the Crabtree effect upon addition of glucose (20) is possible only in intact cells. R in human cells (Table 2) ranges from 4 pmol/s per 106 cells in the small HEK cells (17), 30–40 pmol/s per 106 (HUVEC and fibroblasts; ref. 11, 15), and 70 pmol/s per 106 (mesothelial cells; ref. 19), compared to 250 pmol/s per 106 for the much larger rat hepatocytes (21).

3.1.2. LEAK State

Following stabilization of R, ATP synthesis is inhibited by oligomycin (atractyloside or carboxyatractyloside; Table 1). In this nonphosphorylating or resting LEAK state (L; analogous to State 4; ref. 18), LEAK respiration reflects intrinsic uncoupling as (i) compensation for the proton leak at maximum mitochondrial membrane potential, (ii) proton slip (decoupled respiration), (iii) electron slip which diverts electrons toward reactive oxygen species (ROS) production, and (iv) cation cycling (Ca2+, K+) (3, 4). Monitoring L should be limited to <5 min (Fig. 2), to avoid secondary effects on coupling and respiratory capacity.

3.1.3. ETS Capacity

Mitochondrial respiratory control by the phosphorylation system is partially or fully released by pathophysiological uncoupling and dyscoupling, or experimentally by titration of a protonophore such as FCCP (Table 1). In the noncoupled, open proton circuit, the electrochemical proton potential across the inner mitochondrial membrane is collapsed. Uncoupler titration, therefore, removes

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the electrochemical backpressure on the proton pumps (Complexes CI, CIII, and CIV) and stimulates respiration maximally at level flow as a measure of ETS capacity in the noncoupled state E (Fig. 2). It is important to titrate an optimum concentration of uncoupler, beyond which respiration is inhibited (3, 11). Optimum uncoupler concentrations depend on cell type, cell concentration, medium, and permeabilized versus intact cells. Inhibition of E by oligomycin should be evaluated by uncoupler titrations in the absence of inhibitor. E ranges from 14 to 180 pmol/s per 106 cells in HEK, CEM, HUVEC, fibroblasts, and mesothelial cells, largely depending on cell size. ETS capacity per cell doubles with cell size in CEM cells after cell cycle arrest in the G1- versus S-phase, and in senescent fibroblasts (Table 2). 3.1.4. Residual Oxygen Consumption

Residual oxygen consumption (ROX) remains after inhibition of the ETS. Mitochondrial respiratory states R, L, and E are corrected for ROX (Table 2). Many cellular oxygen consuming enzymes and autooxidation reactions give rise to ROX, including peroxidase and oxidase activities which partially contribute to ROS production. It is difficult to evaluate exactly the extent to which inhibitors of the ETS (Table 1) exert an influence on ROX. Cyanide and azide inhibit CIV and other heme-containing enzymes such as catalase, and may thus modify ROX. Valuable information on ROX is obtained by sequential titration of inhibitors (Fig. 2). Rotenone inhibits cell respiration of human fibroblasts and HUVEC, without a further decline of ROX after the addition of antimycin A (15, 16).

3.1.5. Coupling Control Ratios from the Coupling Control Protocol

Flux control ratios (FCRs) express respiratory control independent of mitochondrial content and cell size. FCRs are normalized for maximum flux in a common reference state, to obtain theoretical lower and upper limits of 0.0 and 1.0 (0 and 100%; Table 2). 1. ROX/E¢: The ROX/E¢ ratio is low (0.01–0.07; Table 2), but ROX contributes to a significant extent to LEAK respiration, with corresponding ROX/L¢ ratios ranging from 0.1 to 0.3, and up to 0.5 in growth-arrested fibroblasts (Table 2). 2. L/E: The LEAK control ratio is the ratio of LEAK respiration and ETS capacity. L/E ranges from 0.09 to 0.14 in various cells (Table 2; the inverse, 11–7, is the respiratory control ratio, RCR; ref. 4, 18). Dyscoupling increases the L/E ratio, e.g., to 0.21 in senescent fibroblasts (Table 2). Alternatively, the L/E ratio may increase without intrinsic uncoupling or dyscoupling, if ETS capacity is diminished. It is, therefore, important to evaluate potential defects of ETS capacity per mt-marker, e.g., ETS per citrate synthase activity (12, 15, 19). 3. R/E: The ROUTINE control ratio is the ratio of (coupled) ROUTINE respiration and (noncoupled) ETS capacity.

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R/E ranges from 0.2 to 0.4 (Table 2; the inverse of 5–2.5 is the uncoupling control ratio, UCR; ref. 3, 11–15). The R/E ratio is an expression of how close ROUTINE respiration operates to ETS capacity. Reported R/E ratios ³ 0.5 (22) could not be reproduced by HRR in a wide range of human cell types and incubation conditions (Table 2). The discrepancies cannot be fully explained by high glucose concentrations in culture and respiration media, since glucose exerts an effect not only on R but also on E (20). R/E ratios increase due to (i) high ATP demand and ADP-stimulated ROUTINE respiration, (ii) dyscoupling (senescent fibroblasts; Table 2), and (iii) limitation of respiratory capacity by the defects of substrate oxidation and complexes of the ETS. 4. (R − L)/E: The netROUTINE control ratio, (R − L)/E, expresses phosphorylation-related respiration (corrected for LEAK respiration) as a fraction of ETS capacity. 0.1–0.3 of ETS capacity is used for oxidative phosphorylation under ROUTINE conditions (Table 2). (R − L)/E remains constant, if dyscoupling is fully compensated by an increase of ROUTINE respiration and a constant rate of oxidative phosphorylation is maintained (fibroblasts in Table 2). Upon stimulation of ROUTINE respiration by an increased ATP demand or if the ETS capacity declines without effect on R, however, (R − L)/E increases, which indicates that a higher proportion of the maximum capacity is activated to drive ATP synthesis. (R − L)/E declines to zero in either fully uncoupled cells (R = L = E) or in cells under metabolic arrest (R = L < E). 5. If the PC protocol is extended by the measurement of cytochrome c oxidase, then the ratio of CIV activity and noncoupled respiration is an index of the apparent excess capacity of this enzyme step in the ETS. Autooxidation of ascorbate and TMPD (Table 1) is extremely high in culture media, hence a mitochondrial respiration medium is used (12). 3.1.6. Respirometric Viability Index

Two quantitative indices of cell membrane permeability (cell viability) are derived from the protocol shown in Fig. 2 (19, 23). (i) The stimulatory effect of succinate + ADP is related to cell membrane permeabilization as (RS − R)/(ES − R). The permeable cells are depleted of substrates and adenylates, hence succinate + ADP stimulates respiration, RS − R > 0, in permeabilized cells only. (ii) Stepwise inhibition by rotenone (CI) and antimycin A (CIII) is related to cell membrane permeabilization as (ROXS − ROX)/ES. ES = ES¢ − ROX is the ETS capacity. CII respiration is inactivated in intact cells after inhibition of CI by rotenone, since external succinate cannot penetrate the plasma membrane, succinate production

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by the tricarboxylic acid cycle (TCA) is stopped when NADH cannot be oxidized, and there are no cytosolic sources of succinate. Addition of succinate to intact cells, therefore, does not exert any effect on respiration, ROXS − ROX = 0 (11, 13). ROXS − ROX represents succinate-supported respiration of permeable cells (in the presence of the CI inhibitor rotenone, Rot) over antimycin A-inhibited ROX of all cells. These respiratory viability indices are based on preserved respiratory function in mt-respiration medium after cell membrane injury, whereas respiration of permeabilized cells is fully inhibited by high Ca2+ in culture media (23, 24). The respirometric approach was confirmed by agreement between respirometric viability (0.79 ± 0.03 and 0.80 ± 0.03 for controls and IL-1b exposed cells) and cell viability (0.87 ± 0.03 and 0.79 ± 0.02; mean ± SD) obtained with a CASY 1 Cell Counter and Analyser System (Schaerfe System, Germany; ref. 19). 3.2. Preparation of Permeabilized Cells and Muscle Fibers

3.2.1. Permeabilization of Cells

Extended functional OXPHOS analysis requires isolation of mitochondria or controlled plasma membrane permeabilization, with effective wash-out of free cytosolic molecules including adenylates, substrates, and cytosolic enzymes, making externally added compounds accessible to the mitochondria (14, 25–29). Full mechanical permeabilization of cell membranes is achieved in liver tissue (29). Biopsy sampling and mechanical fiber preparation lead to partial permeabilization of skeletal muscle. Without homogenous cell membrane integrity, respiration cannot be studied in the ROUTINE state. At low concentrations, digitonin or saponin permeabilize the plasma membranes completely and selectively due to their high cholesterol content, whereas mitochondrial membranes with lower cholesterol content are affected only at higher concentrations. Mitochondrial isolation is more time-consuming than cell membrane permeabilization. Merely 1 or 2 mg wet weight of cardiac or skeletal muscle fibers is sufficient for individual experimental runs with the OROBOROS Oxygraph-2k, but >70 mg is required even for micropreparations of isolated mitochondria (30). The homogeneous suspension of isolated mitochondria yields a representative average for large tissue samples, whereas tissue heterogeneity contributes to the variability of results with small samples of permeabilized fibers. Mitochondria can be isolated to separate different mitochondrial subpopulations (31). Isolated mitochondria and small cultured cells are the appropriate models for the study of mitochondrial oxygen kinetics ((1, 2, 5, 10, 11, 17, 20, 32–34); see Note 2). 1. After air calibration of the oxygen sensors in the O2k-chamber with MiR06, the medium is siphoned off.

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2. Cells are suspended in respiration medium at a final cell density such that ROUTINE respiration yields a volume-specific oxygen flux of about 20 pmol/s per cm3, or higher (Fig. 2). Up to 3 ml are added into each O2k-chamber, while rotation of the stirrers is maintained. Samples can now be collected from the stirred chambers containing a homogenous cell suspension, for the analysis of cell count, cell volume, and cell viability (CASY, Schaerfe System, Germany), protein concentration, and enzyme assays. A minimum of 2.1-ml cell suspension must remain in the chamber. 3. Close the chambers by fully inserting the stoppers into the volume-calibrated position, thereby extruding all gas bubbles. Siphon off any excess cell suspension from the receptacle of the stoppers. 4. Start data acquisition with a new file, and allow endogenous R to stabilize for 15–20 min (compare Fig. 2). 5. Titrate the first selected substrates into the chamber (e.g., glutamate + malate), and add digitonin at optimum concentration, e.g., 10 mg per 106 cells. Observe a gradual decline of respiration due to cell membrane permeabilization and loss of adenylates from the cytosol, allow L to stabilize, and proceed with a SUIT protocol (Subheading 3.3.4). 6. Optimum digitonin concentrations for complete plasma membrane permeabilization of cultured cells can be determined directly in a respirometric protocol (Fig. 3), which may be used simultaneously for selecting optimum experimental cell concentrations. After inhibition of endogenous R by rotenone, respiration of intact cells (viability >0.95) is not stimulated by the addition of succinate and ADP (compare Fig. 2, with cell viability of 0.8). Subsequent stepwise digitonin titration yields gradual permeabilization of plasma membranes, shown by the increase of respiration up to full permeabilization (Fig. 3; at 10 mg per 106 cells). Respiration is constant over a range of optimum digitonin concentrations, but is inhibited at higher concentrations when the outer mitochondrial membrane becomes affected and cytochrome c is released (cytochrome c test, see Subheading 3.3.2). Respiration of permeabilized cells is stable during SUITs in the presence of an optimum digitonin concentration. 3.2.2. Muscle Biopsy and Short-Term Storage

This project was approved by the Ethics Committee of the Medical University of Innsbruck (AN3433 271/4.12). In sports physiology, it is important to choose a muscle for biopsy sampling that is actively involved in the motion of interest. The human muscle most extensively studied for functional diagnosis of mitochondrial

D. Pesta and E. Gnaiger 30 Oxygen flow, IO2 [pmol·s−1·10−6 cells]

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20

10 Succinate+Rotenone+ADP

0 0

10

20

30

Digitonin [µg·cm−3]

Fig. 3. Protocol for determination of optimum digitonin concentration for selective cell membrane permeabilization by respirometry and titration of digitonin to initially intact cells suspended in mitochondrial medium, in the presence of 10 mM succinate, 0.5 mM rotenone, and 1 mM ADP. Human umbilical vein endothelial cells transformed by lung carcinoma at a density of 1.02 (±0.16) × 106 per cm3 (N = 6; ±SD). 12–14 min time intervals between titrations up to 3 mg/cm3, 4–5 min at higher digitonin concentrations. Permeabilization at a digitonin concentration of 10 mg per 106 cells is optimum for ADP-stimulated respiration. From ref. 26.

diseases is the quadriceps (m. vastus lateralis; ref. 4) as it is easily accessible and major nerves and blood vessels lie close to the femur and are unlikely to be injured during biopsy sampling (35). Ultrasound imaging is a simple and quick technique to assess the site and depth of biopsies, can be applied before taking a biopsy, and can be performed in the outpatient clinic. If the depth is not controlled, this may affect the outcome due to differences in fiber type distribution in the muscle (35). Eight O2k-chambers were operated simultaneously for high throughput of biopsied samples. 1. When applying the local anesthetic, it is important not to infiltrate the muscle since several anesthetics exert a direct effect on mitochondrial function. 2. After taking the biopsy, the tissue is removed from the Bergstrom needle with a pair of forceps (rounded tip). The tissue is placed onto a precooled Petri dish on ice, and – if necessary – cut diagonally into small portions of 5–10 mg wet weight (Ww), using a sharp scalpel. These subsamples thus have largely equivalent fiber types. 3. Subsamples of 5 mg Ww for HRR are placed quickly into small tubes with 3–4 ml ice-cold BIOPS for two replicate measurements. In this preservation solution, the sample can be stored for several hours at 0 °C, depending on the source of muscle

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Fig. 4. Preparation of permeabilized muscle fibers from a small biopsy of human vastus lateralis. (a) A large amount of connective tissue (circle) is removed. (b) Muscle fiber bundle. (c) Fiber bundles after mechanical separation with a pair of forceps with very sharp angular tips (standardized period of 4 min). Mesh-like structure and change in color from reddish to pale due to loss of myoglobin and removal of remaining vessels. (d) 12-well plate (Falcon 35/3043) for permeabilization of muscle fibers for four respiratory chambers with sequential incubation in BIOPS, saponin, and MiR06.

tissue (human skeletal muscle for 24 h; ref. 9). This provides the possibility for shipping of biopsies on ice for functional analysis by HRR. 4. Subsamples for other assays (histochemical/morphological; enzymatic; mtDNA) are stored in assay-specific media or liquid nitrogen. 3.2.3. Mechanical Preparation of Permeabilized Fibers

1. The tissue sample with BIOPS solution is transferred onto a small Petri dish on an ice-cold metal plate. 2. Connective tissue is removed using two pairs of very sharp angular forceps (Fig. 4a). 3. Fiber bundles are separated mechanically with these forceps over a standardized period of 4 min for the preparation of a 2-mg sample of human v. lateralis. Fibers are partially teased apart and stretched out, remaining connected in a mesh-like framework (Fig. 4c). Proper separation and a change from red to pale color are best observed against a dark background (Fig. 4a–c). At least during a start-up period, it is recommended to use a dissecting scope for effective removal of connective tissue and observation of the mechanical separation.

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Initially, difficulties arise frequently from the application of excess tissue, which makes mechanical separation tedious. 4. Fiber bundles of similar mass are placed sequentially into 2-ml ice-cold BIOPS in individual wells (Fig. 4d). 3.2.4. Chemical Permeabilization and Wet Weight of Muscle Fibers

1. After fibers for all simultaneously operated O2k-chambers are mechanically prepared and placed into the wells with ice-cold BIOPS, the fiber bundles are transferred quickly into 2 ml freshly prepared saponin solution (50 mg/ml BIOPS; add 20 ml saponin stock solution of 5 mg saponin/ml BIOPS into 2 ml BIOPS; Fig. 4d). 2. Shake by gentle agitation in the cold room (on ice) for 30 min. 3. Transfer all samples from the saponin solution into 2 ml of MiR06 (Fig. 4d). Continue shaking by gentle agitation for 10 min in the cold room (on ice). 4. Wet weight measurements are made after permeabilization, which reduces osmotic variations in water contents. Loosely connected fiber bundles (skeletal muscle 1–3 mg Ww; heart 0.5–2 mg Ww) are taken with the pair of sharp forceps (rounded tip) and placed for 5 s onto dry filter paper. During this time, wipe off any liquid from the tip of the forceps with another filter paper. Take the sample from the filter paper, touch it once more shortly onto a dry area of filter paper while holding it with the forceps, and place the sample onto a small plastic plate on the table of the tared balance. 5. Immediately after reading the Ww, the sample is transferred into a separate droplet of ice-cold MiR06 on a large Petri dish. Check that the tare balance reading returns to zero. Each droplet contains a sample for a respirometric experiment. 6. A pair of forceps with straight tips is used to fully immerse the fibers into the medium in the O2k-chamber. Check if the entire tissue sample has been removed from the droplet into the O2kchamber. 7. Full permeabilization is validated by a decline of LDH activity to 1% of intact tissue (8), or more quickly by respirometry. Respiration of fully permeabilized tissue is not increased by titration of saponin or digitonin in the presence of substrate and ADP. A stimulatory effect of saponin, however, indicates incomplete permeabilization of muscle fibers that were not incubated in saponin solution prior to the experimental run (Fig. 5). Saponin-permeabilization in the respiration chamber does not yield maximum respiratory capacity of muscle fibers. The larger saponin-stimulation of respiration indicated a lower degree of permeabilization (Fig. 5b), which correlated with lower mass-specific oxygen flux even after addition of saponin (compare Fig. 5a).

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Fig. 5. Saponin test for cell membrane permeabilization. Oxygraph traces of oxygen concentration ( cO2 ) and mass-specific oxygen flux [ J O2 (pmol O2/s per mg Ww)]. Fibers of human vastus lateralis were mechanically but not chemically permeabilized before incubation. Partial permeabilization of the cell membrane is shown by the stimulatory effect of glutamate (G) and succinate (S) in the presence of malate, octanoylcarnitine, and ADP (2.5 mM), whereas lack of full permeabilization is seen by further stimulation of respiration after addition of saponin (50 mg/ml). 37 °C, MiR06, 2 ml O2k-chamber. (a) 2.9 mg Ww. (b) 2.7 mg Ww. Experiment 2010-03-08 AB-01.

3.3. HRR with Permeabilized Muscle Fibers, Permeabilized Cells, and Isolated Mitochondria 3.3.1. Temperature

3.3.2. Substrates: Electron Donors

The further the experimental conditions differ from the physiological reference state, the larger the error becomes which may result from adjustment to 37 °C of respiratory fluxes in various metabolic states, applying a commonly assumed constant temperature coefficient. Assuming a Q 10 of 2 (multiplication factor for flux at a 10 °C difference), the temperature coefficients for rates measured at 22, 25, or 30 °C are 2.83, 2.30, and 1.62, respectively, to convert to respiration at 37 °C. Some fundamental functional properties of mitochondria change at 25 °C, for instance, there is a shift from proton leak at 37 °C (proton flux through the membrane) to proton slip at 25 °C (protons pulled back into the matrix phase within a proton pump; ref. 36). In mitochondrial physiology, therefore, experimental temperature close to body temperature has become a standard for quantitative evaluation of mitochondrial respiratory function in mammalian cells (Table 2) and tissues (4). Mitochondrial respiration depends on a continuous flow of electron-supplying substrates across the mitochondrial membranes into the matrix space. Many substrates are strong anions that cannot permeate lipid membranes and hence require carriers. Various anion carriers in the inner mitochondrial membrane are involved in the transport of mitochondrial metabolites.

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Their distribution across the mitochondrial membrane varies mainly with DpH and not with Dy, since most carriers (but not the glutamate-aspartate carrier) operate nonelectrogenically by anion exchange or co-transport of protons. Depending on the concentration gradients, these carriers also allow for the transport of mitochondrial metabolites from the matrix into the cytosol and for the loss of intermediary metabolites into the incubation medium. Export of intermediates of the TCA plays an important metabolic role in the intact cell. This must be considered when interpreting the effect on respiration of specific substrates used in studies of permeabilized cells and isolated mitochondria (4, 37). Some typical saturating substrate concentrations used in respiratory studies are listed in Table 1. 1. Electron donors for NAD+ and Complex I: Substrate combinations of pyruvate + malate (PM) and glutamate + malate (GM) activate dehydrogenases yielding reduced nicotinamide adenine dinucleotide (NADH), which feeds electrons into CI (NADHubiquinone oxidoreductase) and hence down the thermodynamic cascade through the Q-cycle, CIII, cytochrome c, CIV, and ultimately O2. Electrons flow from NADH to oxygen with three proton pumps (CI, CIII, and CIV) in series. 2. Complex II is the only membrane-bound enzyme in the TCA cycle. The flavoprotein succinate dehydrogenase is the largest polypeptide of CII. Following succinate oxidation, the enzyme transfers electrons directly to the quinone pool (38). Whereas CI is NADH-linked upstream to the dehydrogenases of the TCA, CII is FADH2-linked downstream with subsequent electron flow to the Q-junction (4, 37). Electrons flow from succinate to oxygen with two proton pumps (CIII and CIV) in series. 3. Studies of fatty acid oxidation involve a large variety of substrates, such as palmitic acid, palmitoylcarnitine, or palmitoyl-CoA with carnitine. Like CII, electron-transferring flavoprotein (ETF) is located on the matrix face of the inner mitochondrial membrane. It supplies electrons from fatty acid b-oxidation to CoQ. For b-oxidation to proceed, convergent electron flow into the Q-junction is obligatory from both CI and ETF. Malate is provided, therefore, simultaneously with fatty acid substrates, and fatty acid oxidation is blocked by inhibition of CI. Concentrations of fatty acid substrates must be optimized carefully, to reach substrate saturation without inducing inhibitory and uncoupling effects. The octanoylcarnitine concentration was 0.2 mM (39) in the present protocol. Higher concentrations did not yield higher flux. 4. Ascorbate and TMPD (Table 1) are artificial electron donors reducing cytochrome c. Ascorbate is added first, to maintain TMPD in a reduced state. CIII is inhibited by antimycin A or myxothiazol, to activate cytochrome c oxidase (CIV) as

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an isolated step. Autooxidation of ascorbate and TMPD depends on (i) their concentrations, (ii) oxygen concentration, (iii) concentration of added cytochrome c, and (iv) the medium. Histidine stimulates autooxidation of ascorbate and is therefore omitted from MiR06. Chemical background oxygen flux plus ROX is determined at the end of an experimental run after inhibition of CIV by cyanide or azide at low oxygen concentration, continued after reoxygenation at high oxygen concentration. The keto acids pyruvate and a-ketoglutarate remove cyanide from CIV, forming the respective cyanohydrins. Reversibility of cyanide inhibition is particularly effective at high oxygen levels (40). Cyanide cannot be used, therefore, in the presence of pyruvate as a CIV inhibitor. High azide concentrations must be applied for full inhibition of CIV (Table 1). To separate autooxidation from ROX, oxygen consumption is determined in the absence of biological material under experimental conditions as a function of oxygen concentration. The chemical and instrumental background is subtracted from total measured oxygen flux to obtain CIV activity. 5. Cytochrome c does not pass the intact outer mt-membrane. Comparable to the succinate test for plasma membrane permeability, a cytochrome c test can be applied to evaluate the intactness of the outer mt-membrane in mitochondrial preparations (28). Permeabilized fibers of human v. lateralis (healthy controls) do not show any cytochrome c effect when 10 mM cytochrome c is added (Fig. 6). Cytochrome c is added early in the protocol (after ADP; ref. 41) to obtain all active fluxes in a comparable c-activated state, or at a late active state (Fig. 6; or after 100-min incubation; ref. 42), indicating stability of the outer mt-membrane. The kinetic response to external oxidized cytochrome c is monophasic hyperbolic and identical in cytochrome c-depleted permeabilized fibers and isolated mitochondria of rat heart (treated by hypo-osmotic shock), with a c50 of 0.4 mM cytochrome c supporting halfmaximum flux with succinate + rotenone and saturating ADP (28, 33). 10 mM cytochrome c, therefore, is sufficient to saturate electron transfer from CII. In the presence of 0.5 mM TMPD and 2 mM ascorbate, the kinetic response to cytochrome c is biphasic, with a high-affinity K m¢ of 0.5 and 0.9 mM in isolated mitochondria versus permeabilized fibers, and a low-affinity K m¢ of 12 mM in both preparations. Then 10 mM cytochrome c saturates the velocity of CIV to only 63 and 75% in fibers and mitochondria, respectively (28, 33). 3.3.3. ADP and Inorganic Phosphate

ADP and inorganic phosphate are added to permeabilized cells and fibers at high concentrations to saturate OXPHOS capacity. The transmembrane proton pumps drive H+ out of the matrix

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Fig. 6. Oxygen concentration ( cO ) and volume-specific oxygen flux [ J (pmol O2 s−1 cm−3)], as a function of time, in a 2 O2 SUIT protocol with permeabilized human muscle fibers, with octanoylcarnitine + malate and 2.5 mM ADP, addition of glutamate (G), succinate (S), cytochrome c (Cyt c), FCCP (F), and rotenone (Rot; see Table 1). 2.5 and 5 mM ADP resulted in J D2.5 / J D5 flux ratios of 0.86 for CI respiration (a) and CI + II respiration (b); J D2.5 / J E was 0.71 and J D5 / J E was 0.82. J D5 / J E provided an estimate of the phosphorylation system control ratio (P/E ). P/E was 0.80 and 0.82 with CI + II + ETF substrates (a, b). Corresponding ETS capacities at 0.75 and 1.0 mM FCCP (F; three titrations) were (a) JE = 98 pmol O2 s−1 mg−1 Ww (2.2 mg Ww) and (b) 91 pmol O2 s−1 mg−1 Ww (1.8 mg Ww). The additive effect CI + II electron-input is seen by the increase in flux after the addition of succinate and inhibition by rotenone. Reoxygenations by injections of H2O2 into MiR06. 37 °C, 2-ml chamber. Experiments 2010-07-12H-01 and 2010-06-24 H-01.

phase against an electrochemical backpressure, which is used in turn to fuel phosphorylation of ADP and release of ATP at the ATP synthase. The proton circuit is partially coupled in the OXPHOS state, since a fraction of the electrochemical gradient is dissipated through proton leaks. Diffusion restriction as shown by oxygen kinetics (Subheading 3.3.6) and the outer mt-membrane generate barriers for inorganic phosphate and ADP different from isolated mitochondria (2, 8, 27, 43). MiR06 contains 10 mM phosphate. Saturation by ADP requires testing by titrations. At a high apparent Km for ADP of 0.5 mM (27), flux at 2.5 and 5 mM

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ADP is ADP-limited by 13 and 7% (assuming L/P = 0.2). 2.5 mM ADP is saturating in many cases, yet a further increase of ADP concentration provides a test for saturating [ADP]. This is particularly important for the evaluation of OXPHOS versus ETS capacity (P versus E; Fig. 6). 3.3.4. SUIT Protocols

Substrate combinations that match physiological intracellular conditions are applied for the evaluation of coupling control, OXPHOS, and ETS capacities. A new perspective of mitochondrial physiology and respiratory control emerged from a series of studies based on HRR with novel SUIT protocols (37, 41, 44). A SUIT protocol is shown in Fig. 7a, which may be summarized in abbreviated form. Abbreviations and sites of action are listed in Table 1. Subscripts indicate coupling control states (N: no adenylates, D2.5 and D5: 2.5 and 5 mM ADP; F=1.25: stepwise titration for evaluation of optimum 1.25 mM FCCP concentration, inducing the noncoupled state E); ETS inhibitors are in parentheses:

OctM N + D 2.5 + G D2.5 + S D2.5 + D5 + F=1.25 + (Rot )E + (Mna + Myx + Ama )ROX (1) 1. OctMN: (ETF + CI)L, octanoylcarnitine (Oct) and malate (M) in the LEAK state L, in the absence of ADP (no adenylates, N). 2. OctM D2.5 : (ETF + CI)P , OXPHOS (P; State 3) after titration of 2.5 mM ADP (D), flux increases to active respiration, limited by substrate supply to ETF and CI. 3. GMOct D2.5 : (CI + ETF )D2.5 , stimulation by glutamate, with malate as substrates for CI; OXPHOS is ADP-limited at 2.5 mM. 4. GMSOct D2.5 : (CI + II + ETF )D , respiration is further stimu2.5 lated by adding succinate, activating convergent electron flow from CI + II into the Q-cycle (4, 37). 5. GMSOct D5 : (CI + II + ETF)P , OXPHOS capacity at 5 mM ADP. 6. GMSOctE: (CI + II + ETF)E , ETS capacity after FCCP titration in steps of 0.5, 1.0, and 1.25 mM optimal concentration (noncoupled state). Activation by uncoupling is expected if the phosphorylation system (ANT, ATP synthase, phosphate transporter) limits OXPHOS capacity (4, 37), but also if ADP is not saturating. 7. S(Rot)E: CIIE , after inhibition of CI by rotenone, ETS capacity is measured with the entry of electrons from CII only into the Q-cycle. 8. (Mna + Myx + Ama)ROX: ROX is determined after sequential inhibition of the ETS by malonic acid (CII), myxothiazol, and antimycin A (CIII). As shown in the coupling/substrate control diagram (Fig. 7c), alternative protocols are required with separate incubations to obtain information on additional coupling/substrate states. Protocol (2) starts again in state (ETF + CI)L, then places emphasis

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Fig. 7. Substrate–uncoupler–inhibitor titration (SUIT) protocol with substrates for electron-transferring flavoprotein (ETF), Complex I (CI), and Complex II (CII). See also Fig. 5. (a, b) Superimposed oxygraph traces from parallel measurements in two chambers with permeabilized fibers from a biopsy of human vastus lateralis, 3.4 and 2.8 mg Ww. (a) Mass-specific oxygen flux (pmol O2 s−1 mg−1 Ww). (b) Oxygen concentration (mmol/dm3). Oxygen flux is not shown during a few minutes after oxygenations by oxygen or hydrogen peroxide, when the calculation of slopes is disturbed by the step changes in oxygen concentration. 37 °C, MiR06, 2-ml chamber. Experiment 2010-03-04 CD-01. (c) Coupling/substrate control diagram with flux control ratios (FCRs) normalized relative to ETS capacity with convergent CI + II electron input (arrows 1, corresponding to Fig. 6a; FCRs in parentheses are pseudo-state P * at 2.5 mM ADP). Additional protocols (arrows 2 and 3 ) are required to fill in the dashed coupling/substrate states, including overlapping respiratory state S (Rot)E in all cases. Residual oxygen consumption (ROX) is determined as a common step in the three protocols on integrated pathways.

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on coupling control with CI substrates from (CI + ETF)L, (CI + ETF)P, to (CI + ETF)E, and finally on substrate control with (CI + II + ETF)E and CIIE. A cytochrome c test (+c; Fig. 6) is added: OctM N + G N + DP + c P + FE + S E + (Rot )E + (Mna + Myx + Ama )ROX

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For illustration, measurements are added of CIV activity (+AsTm) and chemical background (ROX¢ includes autooxidation after inhibition by KCN; Table 1). In the design of multiple, complementary protocols, it is important to include one or several overlapping coupling/substrate states, providing a quantitative link between the separate experimental incubations (Fig. 7c). 3.3.5. Oxygen Flux, Normalization of Flux, and Flux Control Ratios

Mass-specific flux of permeabilized muscle fibers is expressed per mg wet weight (Fig. 7a), integrating mitochondrial quality and quantity (mt-density). Rather than tabulating mitochondrial respiration in an unnecessary variety of units, SI units provide a standard for expressing oxygen flow (mol O2 s−1; pmol/s) (Table 2) and massspecific oxygen flux (pmol/s per mg). Multiply “bioenergetic” units (ng atom O min−1) by 8.33 to convert to SI units (pmol O2 s−1). To separate the effects of mt-quality from mt-density, a common mt-marker is used for normalization, such as mtDNA (41), citrate synthase activity or CIV activity (12), or cytochrome aa3 content (8). Subsamples or the entire contents can be collected from the O2k-chamber for the analysis of CS activity (29). Respiratory flux control ratios, FCR (Fig. 7c), however, are internal ratios within an experimental run and thus minimize several experimental errors, providing the most powerful normalization of flux (4). Substrate control ratios are FCR at constant coupling state, whereas coupling control ratios are FCR at constant substrate state, relating L and P to E (Fig. 7c). 1. The LEAK control ratio, L/E, expresses uncoupling or dyscoupling, provided that specific limitations of flux by E are considered. L/E increases with uncoupling from a theoretical minimum of 0.0 for a fully coupled system to 1.0 for a fully dyscoupled state. 2. The phosphorylation system control ratio, P/E, increases from a minimum of L/E if the capacity of the phosphorylation system is zero to the maximum of 1.0 if the capacity of the phosphorylation system fully matches the ETS capacity (or in

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fully dyscoupled mitochondria), when there is no limitation of P by the phosphorylation system or the proton backpressure. It is important to separate the effect of ADP limitation from limitation by enzymatic capacity at saturating ADP concentration (Fig. 6). In Fig. 7, the P/E flux ratio was 0.83 at 5 mM ADP, whereas the pseudo-state P control ratio (P*/E) was 0.70 at 2.5 mM ADP, due to ADP limitation of respiration. 3. The conventional respiratory control ratio, RCR (State 3/State 4 or P/L) increases from 1.0 to infinity from fully dyscoupled to fully coupled mitochondria. But the RCR declines with increasing levels of coupling as a function of the phosphorylation system control ratio, P/E. For mathematical reasons, it is more appropriate to use the inverse RCR, which is the L/P ratio with the theoretical boundaries of 0.0 at tight coupling, to the maximum of (L/E)/(P/E) £ 1.0, which becomes 1.0 in fully dyscoupled mitochondria. The RCR is useful only in the limiting case when the P/E ratio is 1.0 (since then L/P = L/E; ref. 4). 3.3.6. Oxygen

Physiological intracellular oxygen levels are significantly lower than air saturation under normoxia, hence respiratory measurements carried out at air saturation are effectively hyperoxic for cultured cells and isolated mitochondria (10). Respiratory capacity, however, of mitochondrial preparations must be evaluated at kinetic oxygen saturation as a reference state. The apparent Km for oxygen (c50 [mM] or p50 (kPa) is controlled by diffusion gradients and metabolic state. Even at 20 mM, oxygen does not limit respiration of isolated mitochondria and small cells (20- to 50-fold above the c50 of 0.4–1.0 mM (32–34)). In permeabilized muscle fiber bundles, however, diffusion restriction increases the sensitivity to oxygen supply to a c50 of 10 mM in the passive LEAK state of respiration, when diffusion gradients are small at low oxygen flux. In the active ADP-stimulated state, the c50 is increased to 40 mM, or 100-fold above that of isolated mitochondria (30 °C; rat soleus and rat heart, incubated with pyruvate and malate (5, 45)). Similarly, oxygen limitation of respiration in permeabilized fibers from human v. lateralis is significant at all levels below air saturation, with a typical c50 of 50 mM for ADP-stimulated respiration (37 °C; Figs. 8 and 9). For the analysis of oxygen kinetics, respiration is monitored during aerobic–anaerobic transitions (Fig. 8a) and plotted as a function of oxygen concentration (Fig. 8b). In many but not all cases, a hyperbolic function provides a good fit (Fig. 8b). Instability of respiration over prolonged periods of incubation time may distort the oxygen kinetics, which is minimized by using larger amounts of tissue in such tests, and evaluated by observation of recovery of oxygen flux after reoxygenation (Fig. 8a). Extrapolation of a hyperbolic relationship below air saturation (Fig. 8b) does not appear to be valid, as seen by the constant flux after reoxygenation (Fig. 8a).

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Multiphasic oxygen kinetics becomes particularly evident in cases when the muscle fiber bundle (Fig. 9a) is mechanically separated further into many small pieces during stirring in the O2k-chamber (Fig. 9b). In the latter case, the oxygen diffusion distance is reduced for an increasing fraction of mitochondria, such that the last phase in the aerobic–anaerobic transition resembles that of isolated mitochondria with a steep decline of flux toward oxygen exhaustion (compare Fig. 9a, b). In summary, elevation of oxygen levels in respirometry with permeabilized muscle fibers is necessary to avoid the development of an artificial hypoxic or anoxic core in the nonperfused fiber bundle (see Note 3). The high degree of oxyconformance in permeabilized

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Fig. 9. Oxygen dependence of respiration in two parallel experiments with samples from a biopsy of human vastus lateralis, 2.6 and 3.1 mg Ww (a, b). The horizontal dotted lines indicate the oxygen concentration of 191 mM corresponding to air saturation at standard pressure (100 kPa; 19.6 kPa partial oxygen pressure). (a) Typical near-hyperbolic oxygen dependence, with an apparent p50 of 5.4 kPa (c50 of 52 mM O2; but nonhyperbolic extension at high oxygen; compare Fig. 8). (b) Multiphasic oxygen dependence in fibers that were disrupted into small pieces by the stirrer in the chamber, without inducing turbidity. The last phase at lowest oxygen levels indicates an apparent p50 equivalent to the kinetics of isolated mitochondria, whereas the first phase at high oxygen levels reflects the oxygen dependence of intact permeabilized fibers. 37 °C, MiR06, 2-ml chamber. Experiment 2010-04-26 GH-01.

fibers is not a kinetic property of the mitochondria, but is largely determined by the geometry of the fiber bundle, with diffusion distances increased from 5–10 mm in cells to >150 mm in the intertwined bundle (5). As a consequence, a compromise is suggested to maintain oxygen levels in the range >250 mM to minimize oxygen limitation of respiration, but <500 mM to avoid extremes of hyperoxia experienced by the peripheral or partially separated mitochondria in the oxygraph chamber. 1. Add 2.2-ml medium into the chamber and insert the stopper incompletely, leaving an air space above the stirred medium. A stopper spacer is used for optimal and reproducible positioning. In this state, the oxygen sensors are air-calibrated (Subheading 3.4.1) while fibers are permeabilized. Remove the stopper, insert a permeabilized fiber bundle into the medium, and insert the stopper incompletely. Inject a few milliliters of oxygen from a gas injection syringe through a needle inserted into the injection port of the stopper and extending into the gas phase, but not into the aqueous phase. Thereby an elevated

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oxygen pressure is created above the stirred aqueous medium. Oxygen in the gas and aqueous phases starts to equilibrate rapidly. When the desired oxygen concentration above 400 mM is nearly reached, close the chamber, thereby removing the gas phase and stopping the equilibration process. 2. Small reoxygenation steps during the experiment can be performed by titrating a few microliters of H2O2 (200 mM in H2O, adjust to pH 6 and keep on ice to minimize autooxidation) into MiR06 (containing catalase). This is sufficient to raise oxygen levels from 250 mM again to 400 mM. After injecting H2O2, the time required for the flux to stabilize depends on the step change of oxygen; smaller steps require less time for stabilization (Figs. 6 and 7). Using H2O2 increases total gas pressure with oxygen pressure. This can generate bubbles in steps from air saturation to >400 mM, that is why O2 gas is applied initially. 3.4. High-Resolution Respirometry 3.4.1. Calibration of the Polarographic Oxygen Sensor

Dissolved oxygen concentration is measured amperometrically by Clark-type polarographic oxygen sensors (POS), containing a gold cathode and Ag/AgCl anode connected electrically by a KCl electrolyte, and separated from the sample by an O2-permeant FEP membrane (0.25 mm). A polarization voltage of 0.8 V is applied to reduce O2 that diffuses from the incubation medium to the cathode through the membrane. O2 is reduced to water, generating a current (hence amperometric) that is linearly proportional to O2 partial pressure, pO2 , in the stirred experimental solution (46). After current-to-voltage conversion and amplification, the raw signal is obtained (1 V/mA; with further amplification by gain settings of 2 or 4). Data sampling at time intervals of 2 s is sufficient for routine applications, 1 s is recommended for kinetic experiments, and it can be reduced to 0.2 s for applications with high sample concentrations. 100 data points are averaged at each data sampling interval. The limit of detection of oxygen concentration extends to 0.005 mM (5 nM) O2. The digital resolution is 2 nM, yielding a 500,000-fold dynamic range up to oxygen saturation. The polarographic oxygen sensors (OROBoPOS) are stable for several months without exchange of membranes or electrolyte (3). A standardized calibration of the linear oxygen sensor includes quality control of signal stability (noise and drift; static two-point sensor calibration) and dynamic calibration of the sensor response time (2). 1. For storage, fill up the clean O2k-chambers completely with 70% ethanol, with the POS and stirrers remaining in the assembled chamber, and the stopper loosely inserted and covered by a lid. Storage in 70% ethanol between experiments can be extended over periods of months, keeping the chamber sterile and the POS immediately ready to use (3). 2. After switching on the instrument, set the experimental temperature, wash with distilled or deionized water while the

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stirrer is on (750 rpm or 12.5 Hz is optimal), and add 2.2-ml experimental medium (MiR06). Insert the stoppers slowly to their volume-calibrated position (2-ml effective volume, plus 0.08 ml to fill the stopper capillary). Siphon off excess medium ejected through the stopper capillary with the integrated suction system (ISS; Fig. 1). 3. Lift the stoppers slightly to introduce an air space above the stirred aqueous medium (open position with stopper spacer), and allow for sufficient time to obtain temperature stability and oxygen equilibration between the gas and aqueous phases. The gas volume has to be exchanged for air, if the medium has not been near air saturation initially, to ensure a well-defined pO2 in the gas phase during air equilibration. Equilibration is a slow process, but stability should be reached within 30–60 min. 4. During this time, a quick stirrer test is performed for dynamic sensor calibration, switching off the stirrer shortly with the consequence of a sharp drop of the POS signal, and observing the exponential increase of the signal after the stirrer is switched on. The corresponding response time is a sensitive indicator of dynamic sensor performance, and deconvolution of the signal is possible for high time resolution in kinetic studies (1, 2, 10). 5. The signal at air saturation provides the first calibration point, with raw signal R1. During this period, the slope of O2 over time must be less than 0.5–1.0 pmol/s per ml, indicating proper signal stability of the POS. Air calibrations are performed daily before starting an experiment (47). 6. Titrate 100 ml freshly prepared 10 mM solution of sodium dithionite to fully exhaust the dissolved oxygen concentration to zero. The zero signal, R0, should be <3% of R1, but most importantly, R0 must be stable (higher stability and lower noise than at air saturation). Occasional checks over a period of months are sufficient (3), except in studies of oxygen kinetics, when shortterm zero drift must be accounted for by internal zero calibration after oxygen depletion by mitochondrial respiration, for resolution in the nanomolar oxygen range (1, 2, 10, 34). 3.4.2. Oxygen Solubility and Concentration

To convert pO2 [kPa] to oxygen concentration cO2 [mM], the oxygen solubility of the medium is calculated as a function of temperature and salt concentration. Oxygen calibration is fully supported by the OROBOROS DatLab software and combines the following information (47): 1. Raw signal [R1 (V)] obtained at air saturation of the medium. 2. Raw signal [R0 (V)] obtained at zero oxygen concentration. 3. Experimental temperature [T (°C)] measured in the thermoregulated copper block (Fig. 1).

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4. Barometric pressure [pb (kPa)] measured by an electronic pressure transducer. 5. The oxygen partial pressure [ pO2 (kPa)] in air saturated with water vapor, as a function of barometric pressure and temperature, calculated by the DatLab software (46). 6. The oxygen solubility [ SO2 (mM/kPa)] in pure water as a function of temperature, calculated by the DatLab software (46). 7. The oxygen solubility factor of the incubation medium (FM) which expresses the effect of the salt concentration on oxygen solubility relative to pure water. In MiR06, FM is 0.92 determined at 30 and 37 °C, and FM is 0.89 in serum at 37 °C. The same factor of 0.89 can be used for various culture media such as RPMI (47). At standard barometric pressure (100 kPa), the oxygen concentration at air saturation is 207.3 mM at 37 °C (19.6 kPa partial oxygen pressure; (46)). In MiR06 and serum, the corresponding saturation concentrations are 191 and 184 mM, respectively. 3.4.3. Oxygen Flux and Instrumental Background

Long-term signal stability and low noise of the oxygen signal are a basis for online display of oxygen flux calculated as the negative time derivative of oxygen concentration. The limit of detection in HRR of oxygen flux is 1 pmol/s per cm3 (0.001 mM/s). With small amounts of sample and correspondingly low respiratory flux per volume, the oxygen capacity of the system provides sufficient time to evaluate the stability of respiratory activity in each metabolic state and to permit complex titration regimes (Fig. 7). At a constant volume-specific flux of 100 pmol/s per cm3, 180 mM O2 is exhausted within 30 min (Fig. 7b). Oxygen consumption by the POS and oxygen diffusion below or above air saturation contribute to instrumental background oxygen flux, which is minimized in HRR and corrected for (1–3). At air saturation, the POS generates a current of about 2 mA at a stoichiometry of four electrons/O2. The O2/electron ratio divided by the Avogadro constant (F = 96,485.53 C/mol) yields the amount of oxygen per Coulomb [1/(4·F) = 2.591 mmol O2 C−1] or O2 flow per current (2.591 pmol/s per mA). At 2 mA per 2 ml, therefore, volume-specific O2 flux or O2 consumption by the POS ( J O2 ,POS ) is 2.6 pmol/s per ml. J O2 ,POS declines to zero as a strictly linear function of pO2 or cO2 under constant experimental conditions. Hence correction for J O2 ,POS is simple and accurate, and does not influence the limit of detection of biological flux. In contrast, the contribution of oxygen diffusion to instrumental background is unpredictable in various respirometric systems, and needs to be determined empirically in the closed chamber in the absence of biological material, as a function of oxygen concentration in the experimental range. Effects of oxygen back-diffusion are minimized

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in the Oxygraph-2k by the large volume (2 ml) and the selection of diffusion-tight materials in contact with the respiration medium: Glass chambers and PVDF or titanium stoppers (not Perspex), magnetic stirrer bars coated by PVDF or PEEK (not Teflon), and Viton O-rings (not silicon). Compared to aqueous media, plastic materials such as Teflon have a >10-fold higher oxygen solubility. Plastic is not feasible for respirometry, since uncontrolled oxygen back-diffusion distorts the respiratory decline of oxygen concentration in a closed chamber. Instrumental background tests were designed to detect and eliminate possible oxygen leaks, introducing this integrated systemic calibration as a key component of quality control in HRR ((1–3, 48); see Note 4). 1. Close the chamber by fully inserting the stoppers after stabilization at air saturation, excluding any gas bubbles. After 10–15 min, observe instrumental background flux (J °1) which is due to J O2 ,POS only. J O2 ,POS is 2.5–3.5 pmol/s per cm3 at air saturation in the 2-ml O2k-chamber at 37 °C. Agreement with the predicted flux validates the instrumental limit of detection of flux. Higher J °1 is due to microbial contamination of the medium and chamber. J °1 would increase to 25 pmol/s per cm3 in a 200-ml chamber. Whereas J O2 ,POS decreases linearly to zero at anoxia, it increases to 8 pmol/s per cm3 at 500 mM O2, but instrumental background flux J O2 does not conform to J O2 ,POS at these oxygen levels. 2a. Lower experimental oxygen concentrations are obtained by stepwise titration of small volumes of freshly prepared 10 mM solution of sodium dithionite (Na2S2O4; 1.7 mg/ml phosphate buffer, pH 8) into MiR06. Standardized four-step background tests ( J O2 at air saturation, 100, 50, and 20 mM O2) can be performed automatically using the programmed OROBOROS Titration-Injection microPump (TIP2k; ref. 48). Alternatively, the chamber is opened intermittently for flushing the gas phase with nitrogen or argon, and closed at reduced oxygen concentration. The near-linear dependence of J °O2 on oxygen concentration extrapolates to zero oxygen concentration with an intercept a° of −1.5 to −2.5 pmol/s per cm3, which is the oxygen back-diffusion per volume of the chamber at zero oxygen concentration. A typical value of the slope b° is 0.025. More negative values of a° indicate an oxygen leak in the system. 2b. In experiments with permeabilized fibers at elevated oxygen levels, instrumental background is measured in the experimental range after increasing the oxygen concentration in MiR06 (see above), and stepwise measurement of J °O2 at four oxygen levels matching the experimental oxygen regime (Fig. 7b: 370,

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330, 300, and 240 mM). The recommended range is 400–250 mM. 3. Plot J °O2 as a function of average cO2 during the period of determining J °O2 .Calculate a° and b° by linear regression (DatLab), J °O2 = a ° + b ° ´ c O2 . 4. Background-corrected volume-specific oxygen consumption [ JV ,O2 (pmol/s per cm3)] is calculated automatically for each data point (DatLab, online) over the entire experimental oxygen range (1–3), J V,O2 = -

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4. Notes 1. OXPHOS protocols presented in this chapter and instrumental standards in HRR address new challenges in mitochondrial respiratory physiology and pathology. 2. Emphasis is placed on intact cells, permeabilized cells, and permeabilized muscle fibers. It has not been shown if isolation of mitochondria involves the selective loss of damaged mitochondria, but in any case all types of mitochondria are accessible experimentally in permeabilized cells and tissues. Respiration of permeabilized skeletal muscle fibers and isolated mitochondria yields comparable results on OXPHOS capacity (4). 3. A recent methodological presentation of respirometry with permeabilized muscle fibers (49) lacks consideration on oxygen limitation summarized previously (45, 50), tabulates respiration of rat liver homogenate at 30 °C (49) which is actually mechanically permeabilized pig liver measured at 37 °C (29), and restricts discussion of protocols to simple substrate supply (separate CI- or CII-electron entry into the ETS), whereas full OXPHOS capacity can be obtained only with physiological CI + II substrate combinations (4, 30, 37, 41, 42, 44). 4. Demands are increasing for quality control, quality assurance, traceability of calibrations, and standardization of protocols for functional mitochondrial diagnosis in biomedical research and clinical applications.

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Acknowledgments This work was supported by OeNB Jubiläumsfond project 13476 and is a contribution to Mitofood COST Action FAO602. We thank Dr. Michael Schocke who was responsible for taking the human biopsies, and Drs. Robert Boushel, Flemming Dela, Steen Larson, Nis Stride, Dan Kane, and Darrel Neufer for advice in the technique of biopsy sampling. References 1. Gnaiger E, Steinlechner-Maran R, Méndez G, Eberl T, Margreiter R (1995) Control of mitochondrial and cellular respiration by oxygen. J Bioenerg Biomembr 27:583–596 2. Gnaiger E (2001) Bioenergetics at low oxygen: dependence of respiration and phosphorylation on oxygen and adenosine diphosphate supply. Respir Physiol 128:277–297 3. Gnaiger E (2008) Polarographic oxygen sensors, the oxygraph and high-resolution respirometry to assess mitochondrial function. In: Dykens JA, Will Y (eds) Mitochondrial dysfunction in drug-induced toxicity. Wiley, New York, pp 327–352 4. Gnaiger E (2009) Capacity of oxidative phosphorylation in human skeletal muscle. New perspectives of mitochondrial physiology. Int J Biochem Cell Biol 41:1837–1845 5. Gnaiger E (2003) Oxygen conformance of cellular respiration: a perspective of mitochondrial physiology. Adv Exp Med Biol 543:39–56 6. Gnaiger E, Kuznetsov AV, Schneeberger S, Seiler R, Brandacher G, Steurer W, Margreiter R (2000) Mitochondria in the cold. In: Heldmaier G, Klingenspor M (eds) Life in the cold. Springer, New York, pp 431–442 7. Fasching M, Renner-Sattler K, Gnaiger E (2010) Mitochondrial respiration medium – MiR06. Mitochondr Physiol Netw 14(13): 1–4. http://www.oroboros.at 8. Veksler VI, Kuznetsov AV, Sharov VG, Kapelko VI, Saks VA (1987) Mitochondrial respiratory parameters in cardiac tissue: a novel method of assessment by using saponin-skinned fibers. Biochim Biophys Acta 892:191–196 9. Skladal D, Sperl W, Schranzhofer R, Krismer M, Gnaiger E, Margreiter R, Gellerich FN (1994) Preservation of mitochondrial functions in human skeletal muscle during storage in high energy preservation solution (HEPS). In: Gnaiger E, Gellerich FN, Wyss M (eds) What is controlling life? vol 3, Modern Trends in Biothermokinetics. Innsbruck University Press, Innsbruck, pp 268–271

10. Gnaiger E, Méndez G, Hand SC (2000) High phosphorylation efficiency and depression of uncoupled respiration in mitochondria under hypoxia. Proc Natl Acad Sci USA 97: 11080–11085 11. Steinlechner-Maran R, Eberl T, Kunc M, Margreiter R, Gnaiger E (1996) Oxygen dependence of respiration in coupled and uncoupled endothelial cells. Am J Physiol 271:C2053–C2061 12. Renner K, Amberger A, Konwalinka G, Kofler R, Gnaiger E (2003) Changes of mitochondrial respiration, mitochondrial content and cell size after induction of apoptosis in leukemia cells. Biochim Biophys Acta 1642:115–123 13. Steinlechner-Maran R, Eberl T, Kunc M, Schröcksnadel H, Margreiter R, Gnaiger E (1997) Respiratory defect as an early event in preservation/reoxygenation injury in endothelial cells. Transplantation 63:136–142 14. Stadlmann S, Rieger G, Amberger A, Kuznetsov AV, Margreiter R, Gnaiger E (2002) H2O2mediated oxidative stress versus cold ischemiareperfusion: mitochondrial respiratory defects in cultured human endothelial cells. Transplantation 74:1800–1803 15. Hütter E, Renner K, Pfister G, Stöckl P, JansenDürr P, Gnaiger E (2004) Senescenceassociated changes in respiration and oxidative phosphorylation in primary human fibroblasts. Biochem J 380:919–928 16. Hütter E, Unterluggauer H, Garedew A, Jansen-Dürr P, Gnaiger E (2006) Highresolution respirometry – a modern tool in aging research. Exp Gerontol 41:103–109 17. Aguirre E, Rodríguez-Juárez F, Bellelli A, Gnaiger E, Cadenas S (2010) Kinetic model of the inhibition of respiration by endogenous nitric oxide in intact cells. Biochim Biophys Acta. doi:10.1016/j.bbabio.2010.01.033 18. Chance B, Williams GR (1955) Respiratory enzymes in oxidative phosphorylation. I. Kinetics of oxygen utilization. J Biol Chem 217: 383–393

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19. Stadlmann S, Renner K, Pollheimer J, Moser PL, Zeimet AG, Offner FA, Gnaiger E (2006) Preserved coupling of oxidative phosphorylation but decreased mitochondrial respiratory capacity in IL-1b treated human peritoneal mesothelial cells. Cell Biochem Biophys 44:179–186 20. Smolková K, Bellance N, Scandurra F, Génot E, Gnaiger E, Plecitá-Hlavatá L, Ježek P, Rossignol R (2010) Mitochondrial bioenergetic adaptations of breast cancer cells to aglycemia and hypoxia. J Bioenerg Biomembr. doi:10.1007/s10863-009-9267-x 21. Jones DP (1986) Intracellular diffusion gradients of O2 and ATP. Am J Physiol 250:C663–C675 22. Villani G, Attardi G (1997) In vivo control of respiration by cytochrome c oxidase in wildtype and mitochondrial DNA mutation-carrying human cells. Proc Natl Acad Sci USA 94:1166–1171 23. Gnaiger E, Rieger G, Kuznetsov A, Fuchs A, Stadlmann S, Lassnig B, Hengster P, Eberl T, Margreiter R (1997) Mitochondrial ischemiareoxygenation injury and plasma membrane integrity in human endothelial cells. Transplant Proc 29:3524–3526 24. Gnaiger E, Kuznetsov AV, Rieger G, Amberger A, Fuchs A, Stadlmann S, Eberl T, Margreiter R (2000) Mitochondrial defects by intracellular calcium overload versus endothelial cold ischemia/reperfusion injury. Transpl Int 13:555–557 25. Vercesi AE, Bernardes CF, Hoffmann ME, Gadelha FR, Docampo R (1991) Digitonin permeabilization does not affect mitochondrial function and allows the determination of the mitochondrial membrane potential of Trypanosoma cruzi in situ. J Biol Chem 266:14431–14434 26. Gnaiger E, Kuznetsov AV, Lassnig B, Fuchs A, Reck M, Renner K, Stadlmann S, Rieger G, Margreiter R (1998) High-resolution respirometry. Optimum permeabilization of the cell membrane by digitonin. In: Larsson C, Påhlman I-L, Gustafsson L (eds) Biothermokinetics in the post genomic era. Chalmers Reproservice, Göteborg, pp 89–95 27. Saks VA, Veksler VI, Kuznetsov AV, Kay L, Sikk P, Tiivel T, Tranqui L, Olivares J, Winkler K, Wiedemann F, Kunz WS (1998) Permeabilised cell and skinned fiber techniques in studies of mitochondrial function in vivo. Mol Cell Biochem 184:81–100 28. Kuznetsov AV, Schneeberger S, Seiler R, Brandacher G, Mark W, Steurer W, Saks V, Usson Y, Margreiter R, Gnaiger E (2004) Mitochondrial defects and heterogeneous cytochrome c release after cardiac cold ischemia and

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reperfusion. Am J Physiol Heart Circ Physiol 286:H1633–H1641 Kuznetsov AV, Strobl D, Ruttmann E, Königsrainer A, Margreiter R, Gnaiger E (2002) Evaluation of mitochondrial respiratory function in small biopsies of liver. Anal Biochem 305:186–194 Rasmussen UF, Rasmussen HN (2000) Human quadriceps muscle mitochondria: a functional characterization. Mol Cell Biochem 208:37–44 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 Gnaiger E, Lassnig B, Kuznetsov AV, Margreiter R (1998) Mitochondrial respiration in the low oxygen environment of the cell: effect of ADP on oxygen kinetics. Biochim Biophys Acta 1365:249–254 Gnaiger E, Kuznetsov AV (2002) Mitochondrial respiration at low levels of oxygen and cytochrome c. Biochem Soc Trans 30:252–258 Scandurra FM, Gnaiger E (2010) Cell respiration under hypoxia: facts and artefacts in mitochondrial oxygen kinetics. Adv Exp Med Biol 662:7–25 Dubowitz V, Sewry CA (2006) Muscle biopsy: a practical approach. Saunders Elsevier, Philadelphia Dufour S, Rousse N, Canioni P, Diolez P (1996) Top-down control analysis of temperature effect on oxidative phosphorylation. Biochem J 314:743–751 Gnaiger E (ed) (2007) Mitochondrial pathways and respiratory control. OROBOROS MiPNet, Innsbruck. http://www.oroboros.at Sun F, Huo X, Zhai Y, Wang A, Xu J, Su D, Bartlam M, Rao Z (2005) Crystal structure of mitochondrial respiratory membrane protein Complex II. Cell 121:1043–1057 Puchowicz MA, Varnes ME, Cohen BH, Friedman NR, Kerr DS, Hoppel CL (2004) Oxidative phosphorylation analysis: assessing the integrated functional activity of human skeletal muscle mitochondria – case studies. Mitochondrion 4:377–385 Delhumeau G, Cruz-Mendoza AM, Lojero CG (1994) Protection of cytochrome c oxidase against cyanide inhibition by pyruvate and a-ketoglutarate: effect of aeration in vitro. Toxicol Appl Pharmacol 126:345–351 Boushel R, Gnaiger E, Schjerling P, Skovbro M, Kraunsøe R, Dela F (2007) Patients with type 2 diabetes have normal mitochondrial function in skeletal muscle. Diabetologia 50:790–796

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42. Gnaiger E, Wright-Paradis C, Sondergaard H et al (2005) High-resolution respirometry in small biopsies of human muscle: correlations with body mass index and age. Mitochondr Physiol Netw 10(9):14–15. http://www.mitophysiology.org/index.php?gnaigere 43. 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 44. Aragonés J, Schneider M, Van Geyte K et al (2008) Deficiency or inhibition of oxygen sensor Phd1 induces hypoxia tolerance by reprogramming basal metabolism. Nat Genet 40:170–180 45. Kuznetsov AV, Lassnig B, Margreiter R, Gnaiger E (1998) Diffusion limitation of oxygen versus ADP in permeabilized muscle fibers. In: Larsson C, Påhlman I-L, Gustafsson L (eds) Biothermo kinetics in the post genomic era. Chalmers Reproservice, Göteborg, pp 273–276

46. Gnaiger E, Forstner H (eds) (1983) Polarographic oxygen sensors. aquatic and physiological applications. Springer, New York 47. Gnaiger E (2010) Oxygen calibration and solubility in experimental media. Mitochondr Physiol Netw 6(3):1–20. http://www.oroboros.at 48. Fasching M, Gnaiger E (2010) Instrumental background correction and accuracy of oxygen flux. Mitochondr Physiol Netw 14(6):1–12. http://www.oroboros.at 49. Kuznetsov AV, Veksler V, Gellerich FN, Saks V, Margreiter R, Kunz WS (2008) Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nat Protoc 3:965–976 50. Gnaiger E, Steinlechner R, Keriel C, Leverve X, Rossi A, Saks V, Sibille B, Kay L, Novel V, Daneshrad Z, Gellerich FN, Eberl T, Skladal D, Sperl W, Margreiter R (1995) Oxygen sensitivity of respiration in endothelial cells, hepatocytes and permeabilized muscle fibers studied by high-resolution respirometry. J Mol Med 73:B39

Chapter 4 High-Throughput Analysis of Mitochondrial Oxygen Consumption James Hynes, Rachel L. Swiss, and Yvonne Will Abstract Interest in the investigation of mitochondrial dysfunction has seen a resurgence over recent years due to the implication of such dysfunction in both drug-induced toxicity and a variety of disease states. Here, we describe a methodology to assist in such investigations whereby the oxygen consumption of isolated mitochondria is assessed in a high-throughput fashion using a phosphorescent oxygen-sensitive probe, standard microtitre plates, and plate reader detection. The protocols provided describe the required isolation procedures, initial assay optimization, and subsequent compound screening. Typical data is also provided illustrating the expected activity levels as well as recommended plate maps and data analysis approaches. Key words: Mitochondria, OXPHOS, Oxygen consumption, Toxicity, Respiration, Polarimetry, Oxygen-sensitive probes, Toxicity, Drug safety testing

1. Introduction Mitochondrial dysfunction is a common mechanism of drug-induced toxicity and has been implicated with a variety of drug classes (1, 2). This has led to a requirement for a high-throughput method of assessing the metabolic implications of drug treatment. Oxygen consumption measurements are favored in this regard as they provide direct information on the activity of oxidative phosporylation and are therefore highly sensitive to perturbations in mitochondrial function. Such measurements allow the identification of compounds that specifically perturb mitochondrial function while also providing information on the mechanisms involved; electron transport chain (ETC), ATPase, and adenine-nucleotide translocator (ANT) inhibitors, for example, will cause a decrease in ADP-activated respiration

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_4, © Springer Science+Business Media, LLC 2012

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while uncouplers cause as an increase in basal respiration. Traditionally, these measurements were performed using standard polarography; however, limited throughput precludes such an approach to this type of application. The necessary throughput is instead achieved using a phosphorescent water-soluble oxygen probe (MitoXpress™) thereby allowing 96- and even 384-well plate-based analysis of mitochondrial oxygen consumption (3, 4). Probe emission is quenched by molecular oxygen via a physical (collisional) mechanism; whereby depletion of dissolved oxygen causes an increase in probe emission. Measuring this signal, therefore, allows the quantification of dissolved oxygen, with changes in probe signal reflecting changes in oxygen concentration within the sample. Successful analysis requires rates of oxygen consumption which exceed the rates of back diffusion from ambient air. For this reason, a sealing layer of mineral oil is applied to limit such back diffusion, thereby increasing assay sensitivity. Detailed protocols are presented describing mitochondrial isolation from relevant tissues and typical activity values are provided. Assay optimization is also addressed and suggestions are provided on how to perform compound screening and on the recommended approach to data interpretation.

2. Materials 2.1. Mitochondrial Isolation

1. Glass tissue homogenizer with Teflon pestle (100 ml). 2. Glass beakers. 3. Glass stirring rods. 4. Plastic funnel. 5. Centrifuge tubes (50 ml). 6. Power drill (hand held or static). 7. Ultra-Turrax tissue homogenizer (IKA, T25). 8. Cheesecloth. 9. Refrigerated high-speed centrifuge. 10. Several ice buckets. 11. BCA kit for protein determination. 12. Eppendorf tubes. 13. Standard clear bottom 96-well plate. 14. Automated Pipettes: Gilson P20, P200, and P1000. 15. Eppendorf syringe dispenser with 2.5-ml plastic syringes. 16. Absorbance plate reader. 17. UV reader such as SpectraMax, Tecan, Victor, FLUOstar Omega. 18. Triton X-100. 19. Type XXIV protease.

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Prepare the following buffers (see Note 1.1): 1. Buffer I: 210 mM mannitol, 70 mM sucrose, 5 mM HEPES, 1 mM EGTA, 0.5% BSA, pH 7.4. 2. Buffer II: 210 mM mannitol, 70 mM sucrose, 10 mM MgCl2, 5 mM K2HPO4, 10 mM MOPS, 1 mM EGTA, pH 7.4.

2.1.2. Isolation of Cardiac and Skeletal Muscle Mitochondria

Prepare the following buffers (see Note 1.1): 1. Buffer I: 100 mM KCl, 40 mM Tris–HCl, 10 mM Tris-base, 5 mM MgCl2, 1 mM EDTA, and 1 mM ATP, pH 7.4. 2. Buffer II: 100 mM KCl, 40 mM Tris–HCl, 10 mM Tris-base, 1 mM MgSO4, 0.1 mM EDTA, 0.2 mM ATP, and 2% BSA, pH 7.4. 3. Buffer III: 100 mM KCl, 40 mM Tris–HCl, 10 mM Tris-base, 1 mM MgSO4, 0.1 mM EDTA, and 0.2 mM ATP, pH 7.4. 4. Buffer IV: 220 mM mannitol, 70 mM sucrose, 10 mM Tris– HCl, and 1 mM EGTA, pH 7.4.

2.2. Assay Optimization (see Note 1.1)

1. Isolated mitochondria of known concentration. 2. Ice (for storage of isolated mitochondria). 3. Glutamate/malate: 0.5/0.5 M in H2O, pH 7.4, aliquot and store at −20°C. 4. Sodium succinate: 1 M in H2O, pH 7.4, aliquot and store at −20°C. 5. Adenosine 5¢-diphosphate (ADP): 100 mM in H2O, aliquot and store at −20°C. 6. “Respiration Buffer”: 250 mM sucrose, 15 mM KCl, 1 mM EGTA, 5 mM MgCl2, 30 mM K2HPO4, pH 7.4. 7. MitoXpress™ Probe (Luxcel Biosciences). 8. Heavy mineral oil (VWR, IC15013880). 9. Black body clear bottom 96-well plate (Costar 3631 or equivalent). 10. Automated pipettes: Gilson P20, P200, and P1000. 11. 8- or 12-channel 100-ml pipette. 12. Gilson Distriman® pipette with 1,250-ml cartridges. 13. Multio-BlokR heater. 14. 2 ml clear Eppendorf tubes. 15. Water bath, 30°C (for warming solutions). 16. Time-resolved fluorescence plate reader with temperature control and kinetic analysis software.

2.3. Screening

Materials as listed in Subheading 2.1 plus: 1. PCR plates for compound dilutions.

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2.4. Data Analysis

1. Plate reader software [Magellan (Tecan), MARS (BMG Labtech), WorkOut (PerkinElmer), SoftMax Pro (Molecular Devices)]. 2. Data Processing Software (MS Excel, Microcal Origin, GraphPad prism).

3. Methods The most critical aspect of functional mitochondrial measurements such as those outlined here is the quality of the mitochondria preparation used. Quality control of the mitochondrial preparation is therefore of critical importance such that variations in source tissue or possible batch-to-batch variability can be accounted for prior to the assessment of effector action. It is also critically important that the preparation be well coupled. This is determined from the respiratory control ratio (RCR); the ratio of ADPstimulated to basal respiration (state 3/state 2). Screening for compounds with a mitochondrial liability can be carried out on a variety of substrate combinations, the most common of which are glutamate/malate, and succinate feeding complex 1 and complex 2 of the ETC, respectively. If heart mitochondria are used, fatty acids can be used as substrates, also. Respiration can then be assessed in basal (state 2) or ADP-stimulated (state 3) conditions with inhibitor screening generally performed in state 3 and uncoupler screening performed in state 2. As the resultant oxygen consumption rate is dependent on both substrate and ADP availability, it is necessary to establish the optimum protein concentration for such screening. 3.1. Mitochondrial Isolation

1. Euthanize animals with an overdose of carbon dioxide (see Note 1.2), excise organs rapidly, and place in ice-cold Buffer I.

3.1.1. Isolation of Liver Mitochondria

2. Using a pair of scissors, finely mince approximately 6 g of liver tissue and then wash repeatedly in Buffer I until the homogenate is blood-free. Then add five volumes of Buffer I and homogenize using 6–8 passes of a smooth glass grinder with Teflon pestle driven by a power drill on low speed. 3. Adjust to 8 volumes with Buffer I and centrifuge at 700 × g at 4°C for 10 min, then filter through two layers of cheesecloth and recentrifuge for 10 min at 14,000 × g to precipitate the mitochondrial fraction. 4. Discard the supernatant, using a glass stirring rod, resuspend the mitochondrial pellet in 20 ml of isolation buffer I, and recentrifuge at 10,000 × g for 10 min at 4°C. 5. Repeat this wash step in Buffer II.

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6. Resuspend the resultant mitochondrial pellet in 0.7 ml of Buffer II and store on ice until required (see Note 1.3). 7. Determine protein concentration using a BCA kit as per manufacturer’s instructions, briefly. 8. Construct a standard curve using Albumin stock solutions at 1.5, 1.0, 0.75, 0.5, 0.25, 0.125, 0.025, and 0 mg/ml prepared in 1% v/v Triton X (see Note 1.4). 9. Dilute samples of the mitochondrial preparation 1:60, 1:80, and 1:100, using 1% v/v Triton X as dilute. 10. Mix 9.8-ml protein reagent A with 200-ml protein reagent B to produce the developing reagent. Then add 20 ml of each standard or sample into a 96-well plate followed by 200 ml of developing reagent. 11. Incubate the plate for 30 min at 37°C, then read absorbance at 520 nm and calculate mitochondrial protein concentration (see Note 1.3). 3.1.2. Isolation of Cardiac and Skeletal Muscle Mitochondria

1. Euthanize animals with an overdose of carbon dioxide (see Note 1.2), excise either two rat hearts or 10 g of mixed muscle (EDL/gastrocnemius/soleus), and place in ice-cold Buffer I. 2. Add Type XXIV protease to a concentration of 5 mg/g of wet tissue and mince finely using a pair of scissors. Incubate for 7 min intermittently mixing and mincing, then add an equal volume of Buffer I to terminate the digestion. 3. Using an Ultra-Turrax tissue homogenizer (IKA, T25) set at setting 1 (11,000 rpm) homogenize the resultant mixture for 30 s and then centrifuge the homogenate at 4°C for 10 min at 700 × g. 4. Filter the supernatant through two layers of cheesecloth and recentrifuge at 14,000 × g for 10 min at 4°C. 5. Discard the supernatant, and using a glass stirring rod resuspend the mitochondrial pellet in Buffer II, then centrifuge at 7,000 × g for 10 min at 4°C. 6. Discard the supernatant, using a glass stirring rod, resuspend the mitochondrial pellet in 20 ml of Buffer III, then centrifuge at 3,500 × g for 10 min at 4°C. Then resuspend the mitochondrial pellet in a minimal volume of solution IV for further use (see Note 1.3). 7. Protein concentrations should be determined as described in step 8 of the preceding protocol.

3.2. Oxygen Consumption Analysis: Assay Optimization

1. Reconstitute MitoXpress™ probe in 1 ml of respiration buffer mixing to insure resuspension. Then dilute to 10 ml with the same buffer and warm to 30°C (see Note 2.1).

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2. Set instrument temperature to 30°C and allow sufficient time for target temperature to be reached. Prepare a kinetic protocol using the recommended optical settings (see Table 1) reading test wells at 0.5–1.5 min intervals over 30–60 min (see Note 2.2). 3. Warm mineral oil to 30°C. 4. For state 2 respiration (Basal), add 150 ml of substrate stock solution (succinate or glutamate/malate) to 1.35 ml of incubation buffer and warm to 30°C. 5. For state 3 respiration (ADP-stimulated), add 150 ml of substrate stock and 100 ml ADP stock to 1.25 ml of incubation buffer and warm to 30°C. 6. Using respiration buffer, prepare a six-point mitochondrial protein dilution series to a 1.5-ml total volume for each concentration. Recommended concentrations (mg/ml): 1.5, 1.0, 0.5, 0.25, 0.125, and 0.63. 7. Place a clear bottomed, 96-well plate on a plate heater equilibrated to 30°C and, using automatic or multichannel pipette, dispense solutions as per the recommended plate map (Fig. 1) adding the following to each well: (a) 100 ml of assay buffer containing MitoXpress™ probe. (b) 50 ml of mitochondrial stock dilutions giving the desired mitochondria concentration. (c) 50 ml of substrate solution (final concentration 25 mM for succinate or 12.5/12.5 mM glutamate/malate) for basal respiration. (d) 50 ml of substrate/ADP solution (final concentration 25 mM for succinate, 12.5/12.5 for mM glutamate/malate, 1.65 mM for ADP) for ADP-stimulated respiration. 8. Using a syringe dispenser, add 100 ml of prewarmed heavy mineral oil to each well (see Note 3). 9. Insert the microplate into the fluorescence plate reader and read using the settings outlined above. When measurement is completed, remove the plate and save the data to file. 3.3. Oxygen Consumption Analysis: Screening

1. Reconstitute MitoXpress™ probe in 1 ml of assay buffer mixing to insure resuspension. Then dilute to 10 ml with the same buffer and warm to 30°C. 2. Set instrument temperature to 30°C and allow sufficient time for target temperature to be reached. Prepare a kinetic protocol using the recommended optical settings (see Table 1) reading test wells at 0.5–1.5 min intervals over 30–60 min.

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Fig. 1. Assay optimization. (a) Typical plate layout for the initial optimization of liver mitochondrial protein concentration needed for screening of NCEs. Typical data output is presented in (b) as a MARS plate view (BMG Labtech) showing a serial dilution of mitochondrial protein at the indicated concentrations (mg/ml) measuring both glutamate/malate (left ) and succinate (right ) driven respiration in both basal (state 2, top) and ADP-stimulated (state 3, bottom). State 2 succinatedriven respiration profiles are presented in detail in (c) and the effect of ADP addition on succinate-driven respiration is presented in (d). If cardiac mitochondria are used, the same plate layout can be used to optimize the protein amount for respiration using fatty acids.

3. Prepare compounds in DMSO to 100× the required concentration (see Note 2.4): (a) For initial screening assay at a single concentration (typically 100 nmol/mg) in duplicate as per plate map (Fig. 2). (b) For subsequent mechanistic elucidations and dose–response analysis 1:2 dilution seria are prepared for each compound at one data point per individual concentration as per plate map (Fig. 3). 4. Prewarm mineral oil to 30°C. 5. Prepare substrate stocks and prewarm reagents to 30°C. (a) State 2 analysis (uncoupler screening): Mix 5.4 ml of incubation buffer and 600 ml of either succinate or glutamate/ malate stock.

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Table 1 Summary of recommended instrument setting for common fluorescence plate readers Prompt fluorescence

Time-resolved fluorescence

SpectraMax Gemini (Molecular Devices)

Safire/Genios Pro (Tecan)

FLUOstar Omegaa (BMG Labtech)

Victor™ X4 (PerkinElmer)

Light source

Xe-flashlamp

Xe-flashlamp

Xe-flashlamp

Xe-flashlamp

Optical configuration

Monochromator based Top and Bottom reading

Filter based

Filter based

Filter based

Top reading

Top and Bottom reading

Top reading

Excitation (nm)

380

380 ± 20

340 TR-Ex

340 ± 40

Emission (nm)

650

650 ± 20

655 ± 25

642 ± 10

Measurement mode

Prompt fluorescence Analog mode

TRF Photon counting

TRF Analog mode

TRF Photon counting

TRF: delay/gate times (ms) Intensity mode n.a

70/30

70/30

70/30

Ratiometric mode Delay/Gate 1 Delay/Gate 2

n.a. n.a.

n.a. n.a.

30/30 70/30

30/30 70/30

SoftMax Pro

Magellan™

MARS

Work out

Data analysis package a

A specific script is required from manufacturer to allow ratiometric measurements

(b) State 3 analysis (inhibitor screening): Mix 5 ml of incubation buffer and 600 ml of either succinate or glutamate/malate stock 400 ml of ADP. 6. Prepare 6 ml of stock mitochondria at 4× the required protein concentration (optimized above). 7. Place a black 96-well plate on a plate heater equilibrated to 30°C and dispense the following solutions using either an appropriate automatic or multichannel pipette (see Note 2.3): (a) 100 ml of prewarmed assay buffer. (b) 1 ml of compound at desired concentration, according to the appropriate plate map. (c) 50 ml of mitochondria at optimal dilution. (d) 50 ml of prewarmed substrate stock solution to each well. (e) 100 ml of prewarmed heavy mineral oil.

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Fig. 2. Single dose screening. 46 NCEs can be screened in duplicate on one plate using the plate layout presented in (a). Two vehicle controls (DMSO) are included on the left site of the plate, whereas on the right site of the plate, one would include FCCP [for state 2, (b)] or rotenone [for state 3, (c)], as positive controls giving the maximum uncoupling or inhibition. Typically, compounds showing more than 50% change from control values are flagged and taken forward into dose– response analysis.

8. Place the microplate in a fluorescence reader preset as described above and commence reading. When completed, save data to file. 3.4. Data Analysis 3.4.1. General Approach

1. Standard analysis: The standard data analysis approach entails calculating “slope” values from plots of raw intensity or lifetime versus time using standard linear regression (Fig. 1). This can be performed on the software of most readers [MARS (BMG Labtech); Workout (PerkinElmer); Magellan™ (Tecan); SoftMax Pro (Molecular Devices)], or on packages such as MS Excel. 2. Detailed analysis: For quantitative analysis, linearize the probe profiles using the following co-ordinate scale (5): (a) Abscissa, Y: I0/(I0 − It) – where I0 and It represent fluorescence signals at time 0 and t (b) Ordinate, X: 1/t, min−1 (c) Exclude zero time points Then apply linear regression analysis to the transformed profiles and determine the reciprocal slope for each of the transformed profiles (see Note 2.5).

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a

c

b

e

2600 2400 2200 2000 1800 1600 1400 1200 1000 800 600 400

250 200

O2 µM

Intensity (RFU)

d

150 100 50

0

2

4

6

8

10

12

14

16

0

18

0

2

4

6

Time (min)

f

g 1/Onset Time(mine-1)

O2 Consumption

45 40 35 30 25 20 15 1

10

100

[Drug] nmol/mg

10

12

14

16

18

0.36 0.31 0.26 0.21 0.16 1

10

8

Time (min)

10

100

1000

[Drug] nmol/mg

Fig. 3. Dose–response analysis. Typical plate layout for generation for NCE dose–response curves. 14 compounds can be screened at six different concentrations (200, 100, 50, 25, 12.5, and 6.25 nmol/mg protein) using the plate map presented (a). Typical data output for uncouplers and inhibitors are shown in (b) and (c), respectively (Tecan Magellan plate views). For state 2 analysis, an FCCP dose–response is included in H1–H6 and for state 3 respiration, a nefazodone dose–response is included. A vehicle control is included in D7–12. Raw data are analyzed and UC50/IC50 values generated . Sample uncoupler raw data (A7–12), transformed data, and the resultant dose–response curve are presented in (d), (e), and (f), respectively. (g) Dose–response curve (SoftMax Pro, Molecular Devices) generated using the more basic time-to-threshold data analysis approach available on most plate readers (see Note 2.6).

3. High-throughput analysis: Data analysis for large data sets is best performed using one of the following two approaches (see Note 2.6): The first is to convert probe signal to an oxygen scale and subsequently apply linear regression analysis to the transformed profiles (Fig. 3d–f ). Oxygen conversion can be approximated by normalizing profiles (dividing each value in the profile but the first value) and using the following equation: [O 2 ](t ) = 235 ´ 1 ´ ( S - X ) / ( X ( S - 1)), where S is the signal increase on deoxygenations. For a more detailed analysis, see Note 2.6 and supplemental material I.

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The second involves using a “time-to-threshold” value calculated using plate reader software and relating the reciprocal of this value to drug concentration to generate dose–response information. This can be performed on most plate reader software (Fig. 3g). 3.4.2. Assay Optimization

1. Open the saved file and either analyze on plate reader software (preferable) or export to external software package such as MS Excel. Then plot raw intensity or lifetime versus time (Fig. 1). 2. Using the quantitative data analysis approach outlined above calculate an RCR and compare to expected values to insure the mitochondrial preparation is of sufficient quality (see Note 2.7). RCR is calculated by determining the state 3/state 2 ratio using reciprocal slopes. 3. If this value is sufficiently high, examine individual profiles and select mitochondrial protein concentration that produce reliably measurable signal changes to allow analysis of both inhibition and uncoupling. These are then used for subsequent screening (see Notes 2.8 and 2.9).

3.4.3. Screening

1. Open the saved file and either analyze on plate reader software (preferable) or export to an external software package such as MS Excel. Then plot raw intensity or lifetime versus time. 2. Calculate “slope” values from plots of raw intensity or lifetime versus time using standard linear regression and compare calculated slopes to those of the untreated sample to determine what compounds, if any, are inhibiting/uncoupling (Fig. 2). 3. Dose-based compound ranking can also be performed if necessary by calculating an IC50 (inhibitory effect in state 3) or UC50 (uncoupling effect in state 2 using FCCP as 100% uncoupling) value for each compound. This is best performed using one of the approaches outlined above (see Note 2.6, and supplemental data).

4. Notes 1. Mitochondrial isolation Solutions should be prepared using Millipore grade water and stored in prewashed glassware. pH should be adjusted with HCl and KOH. Do NOT use NaOH. All substrate stocks can be prepared in advance and stored in aliquots at −80°C. Buffers should be freshly prepared on a weekly basis and any BSA additions should be made on the day of use.

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Sprague–Dawley rats (Charles River, Wilmington, MA) or equivalent strain. Care and maintenance need to be in accordance with the principles described in the Guide for Care and Use of laboratory Animals (NIH Publication 85–23, 1985) or equivalent. For best results the animals should weigh between 150 and 180 g. Rats are housed in pairs in a controlled environment with constant temperature (21 ± 2°C) and a 12-h light/dark cycle. Food and water are provided at ad libitum. Avoid anesthetics as they can have adverse effects on mitochondrial quality. Mitochondria should be stored on ice at a protein concentration of >30 mg/ml and used within 4–6 h. Protein standards: can be made in batches and stored at 4°C for several weeks. Albumin is provided as a 2-mg/ml stock solution. 2. Oxygen consumption analysis Standard probe package is for one 96-well plate (or ~100 assay points). Probe diluted in respiration buffer should be used on the same day. Adjust volumes accordingly for smaller numbers of samples. If reconstituted in 1-ml H2O probe stock can be stored in the dark at +4°C for several days. Where possible it is recommended that data be generated in “lifetime” mode. This is achieved on certain instruments with timeresolution capability (e.g., PerkinElmer Victor, BMG FLUOstar Omega) using a ratiometirc read as outlined in Table 4.1. Using these dual intensity reads, the corresponding lifetime is calculated using the following relationship: t = 40/ln(R1/R2), where R1 and R2 represent read one and read two, respectively, providing lifetime values with units of ms. This can be conducted on plate reader software or after export to software packages such as MS Excel or Microcal Origin. If lifetime measurements are not possible, data can be generated in standard “intensity” mode. To minimize oxygen depletion in samples prior to the measurement, plate preparation time should be kept to a minimum. Concentration range is defined by the user. As drug concentrations are usually expressed in nmol/mg of mitochondrial protein, starting concentration should be altered based on the mitochondrial protein concentration being used. DMSO content should not exceed 0.5% v/v and dilution plates should be prepared no earlier than the 24-h pretreatment. If a significant baseline drift is seen on the original profiles (i.e., at zero enzyme concentration), it is recommended that the baseline be subtracted from all profiles prior to the transformation. This usually improves the linearity of transformed plots. While profile linearization provides an accurate measure of enzymatic activity (5) applying this approach to large data sets

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can be unwieldy. In instances where IC50/UC50 data is being generated on large data sets to facilitate compound ranking, less sophisticated approaches can be applied to generate comparable data in a more user friendly, high-through put compatible manner. The options are Oxygen conversion and linear regression Converting from probe signal to an oxygen scale allows the use of simple linear regression for the generation of dose– response data. Oxygen concentrations can be estimated from measured probe intensities using the following relationship: [O 2 ](t ) =

[O 2 ]a ´ I a ´ (I o - I t ) I t ´ (I o - I a )

[O2]a is oxygen concentration in air-saturated buffer (235 mM at 30°C) and It, Ia, and Io are fluorescent signal at time t, signal in air-saturated buffer, and signal in deoxygenated buffer (maximal signal), respectively. By normalizing each profile for initial intensity (dividing each profile by the initial read) and measuring the signal increase on complete deoxygenation (S), this can be simplified to: [O 2 ](t ) = 235 ´ 1 ´ ( S - X ) / ( X ¢( S - 1)) This specific transformation is only valid at 30°C and the signal increase on deoxygenation is specific to reader type, measurement settings, and buffer composition. Plates should be at measurement temperature prior the reading. It is also important that no significant deoxygenation has occurred prior to read 1 of the kinetic analysis. It should be noted that this transformation, in this instance, is not primarily intended as an analytical determination of dissolved oxygen but rather as an estimation of oxygen concentration to facilitate simple linear regression analysis to be performed on large data sets. Threshold time By assessing the time at which a particular signal threshold is reached and using the inverse of this value (1/threshold time) as a metric of oxygen consumption rate, dose– response data can be quickly generated using standard plate reader software. Oxygen consumption rates and the optimum threshold should, however, be optimized such that the threshold is not exceeded prior to the beginning of the assay (uncoupler analysis) or that significant activity is not missed (inhibitor analysis).

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The RCR is calculated by assessing the effect of ADP addition on mitochondrial oxygen consumption and is an important metric in assessing the quality of a mitochondria preparation. A lower than expected value indicates that the mitochondria are insufficiently coupled, probably due to membrane damage during preparation. Rat liver mitochondria generally show RCRs of approximately 5.0 for glutamate/malate-driven respiration or approximately 3.0 for succinate-driven respiration. It should be noted, however, that RCR values are tissuedependent. Calculated RCR values should be compared with that of previous experiments or literature data (6). A concentration producing large signal changes without inducing rapid signal saturation is deemed “optimal,” thereby allowing reliable analysis of both uncoupling and inhibition of mitochondrial. These optimal concentrations will be dependent on the tissue of origin, the substrate used, and the availability of ADP specific. Typical values for rat liver mitochondria are as follows: (a) Basal respiration glutamate/malate: 1 mg/ml. (b) ADP-driven respiration 0.25 mg/ml.

with

glutamate/malate:

(c) Basal respiration with succinate 0.5 mg/ml. (d) ADP-driven respiration with succinate: 0.25 mg/ml. Prior to screening for compounds with a mitochondrial liability, it should be insured that assay performance is consistent. This is determined from CV values for intra- and interassay variations. These values should not exceed 15% and normally be at ~10%. Once mitochondrial preparations are seen to be reproducible and the assay is well established (n = 3–5), this optimization becomes unnecessary. References 1. Dykens JA, Marroquin LD, Will Y (2007) Strategies to reduce late-stage drug attrition due to mitochondrial toxicity. Expert Rev Mol Diagn 7:161–175 2. Wallace KB (2008) Mitochondrial off targets of drug therapy. Trends Pharmacol Sci 29:361–366 3. Papkovsky DB, Hynes J, Will Y (2006) Respirometric screening technology for ADMETox studies. Expert Opin Drug Metab Toxicol 2(2):313–323 4. Will Y, Hynes J, Ogurtsov VI, Papkovsky DB (2007) Analysis of mitochondrial function using

phosphorescent oxygen-sensitive probes. Nat Protoc 1:2563–2572 5. Ogurtsov VI, Hynes J, Will Y, Papkovsky DB (2008) Data analysis algorithm for high throughput enzymatic oxygen consumption assays based on quenched-fluorescence detection. Sens Actuat B Chem 129: 581–590 6. Hynes J, Hill R, Papkovsky DB (2006) The use of a fluorescence-based oxygen uptake assay in the analysis of cytotoxicity. Toxicol In Vitro 20:785–792

Chapter 5 Modulation of Cellular Respiration by Endogenously Produced Nitric Oxide in Rat Hippocampal Slices Ana Ledo, R.M. Barbosa, and J. Laranjinha Abstract Nitric oxide (•NO) is a ubiquitous signaling molecule that participates in neuromolecular phenomena associated with memory formation as well as in excitotoxicity. In the hippocampus, neuronal •NO production is coupled to the activation of the NMDA-type of glutamate receptor. More recently, Cytochrome c oxidase has emerged as a novel target for •NO, which competes with O2 for binding to this mitochondrial complex. This reaction establishes •NO not only as a regulator of cellular metabolism but possibly also as a regulator of mitochondrial production of reactive oxygen species which participate in cellular signaling. A major gap in the understanding of •NO bioactivity, namely, in the hippocampus, has been the lack of knowledge of its concentration dynamics. Here, we present a detailed description of the simultaneous recording of •NO and O2 concentration dynamics in rat hippocampal slices. Carbon fiber microelectrodes are fabricated and applied for real-time measurements of both gases in a system close to in vivo models. This approach allows for a better understanding of the current paradigm by which an intricate interplay between •NO and O2 regulates cellular respiration. Key words: Nitric oxide, Oxygen, Hippocampus, Carbon Fiber Microelectrode, Nafion®, o-phenylenediamine

1. Introduction Like oxygen, nitric oxide (•NO) is a small diatomic molecule essential in physiological processes, including cellular respiration. •NO is a hydrophobic gas that diffuses relatively free in the cellular environment, crossing cell membranes unassisted by receptors or channels. In the late 1980s •NO was shown to regulate multiple signaling pathways in the nervous, immune, and endothelial systems (1). The better characterized intracellular molecular target of •NO is soluble guanylate cyclase, an enzyme activated by •NO binding

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_5, © Springer Science+Business Media, LLC 2012

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that then converts GTP to cGMP (2), which, in turn, initiates signaling cascades leading, for instance, to vasodilation and neuromodulation (3, 4). In addition to guanylate cyclase, cytochrome c oxidase (CcO) is gaining ground in being established as a critical target and mediator of •NO bioactivity. CcO is the terminal complex of the mitochondrial respiratory chain; it accepts electrons from cytochrome c and reduces O2 to H2O. Low concentrations of •NO can reversibly bind to CcO and inhibit mitochondrial respiration by competing with O2 (5–8). Both O2 and •NO bind to the active site of CcO with similar binding affinities (~2 × 108 and 0.4 × 108, respectively) (9–12). Several mechanisms have been proposed to explain the molecular details of the inhibition of CcO by •NO. However, due to the difficulties associated with the measurement of •NO binding to CcO in vivo, most of what is known is based on theoretical models (13–15). The concept of •NO inhibition of CcO has been clearly demonstrated in cellular and subcellular preparations, namely, mitochondria and synaptosomes (5–7). Here, we present a method that allows for the simultaneous and real-time recording of the profiles of •NO and O2 in hippocampal slices at physiological O2 concentrations. This approach uses a complex biological preparation that retains the cytoarchitectural organization found in vivo and allows for a better understanding of the mechanisms of •NO regulation of mitochondrial respiration (16). Both, the preservation of the tissue integrity and the physiological O2 tensions are critical issues when considering the interplay of • NO and O2 in modulating CcO activity. First, both gases are diffusible and, therefore, tissue tortuosity might affect diffusion. Second, at high concentrations of O2 (such as atmospheric O2 tension or even at higher tensions in cell culture experiments) •NO autooxidation by O2 may occur, yielding reactive species and diverting their respective bioactivity. The hippocampus is a structure of the temporal lobe in the central nervous system involved in memory formation (17). In this region •NO is produced upon activation of the NMDA-type of glutamate receptor (18, 19) and acts as a neuromodulator in molecular phenomena associated to long-term potentiation and depression of synaptic strength (20–23). In the hippocampus, the neuronal circuit (input > dentate gyrus > CA3 subregion > CA1 subregion > output) is organized in a lamellar fashion, so in transversal sections this trisynaptic loop is maintained (24, 25). Furthermore, such brain slices are electrophysiologically and biochemically operative (26) and hence the great interest in the use of acute hippocampal slices. In the method presented here, both endogenous •NO and O2 are measured electrochemically using carbon fiber microelectrodes. This methodology allows real-time and direct measurement of

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the electroactive species of interest with high temporal and spatial resolution. In the case of •NO, the use of carbon fiber microelectrodes modified with Nafion® and o-phenylenediamine has been established by others and us as convenient probes for ex vivo and in vivo measurements (27–29). The use of these polymer films covering the carbon fiber surface allows for an increase of sensor selectivity, a relevant concern considering the relatively high oxidation potential of •NO and the presence of various potential electroactive interferents present in tissue preparations from the nervous system, such as ascorbic acid, biogenic amines, indoles, and their metabolites. In the case of O2, it is habitual to use platinum microelectrodes due to their well-known catalytic properties for O2 reduction. However, some authors argue that, contrary to platinum electrodes, carbon-based electrodes consume much less O2 from the medium, and thus, there is a lesser risk of the electrode per se affecting O2 distribution in the tissue (30, 31). In the following sections, we provide a detailed description of the steps involved in the use of carbon fiber microelectrodes for the evaluation of how endogenously produced •NO can modulate cellular respiration in hippocampal slices, namely, microelectrode fabrication, evaluation and application in simultaneous electrochemical recordings; preparation and determination of standard/stock saturated •NO solutions as well as preparation and use of acute hippocampal slice.

2. Materials and Solutions Unless otherwise mentioned, all reagents used are of analytical grade. 2.1. Saturated Nitric Oxide Stock Solutions

Because the preparation of saturated •NO stock solutions is not a standard procedure and varies in the literature, it is described in detail in this chapter. All handling of •NO gas should be performed in a fume hood. Reaction of gaseous •NO and atmospheric O2 results in the production toxic nitrogen oxides such as •NO2 and N2O3. The setup should contain only inert materials, such as glass, teflon, and stainless steel tubing and fittings. Solvents should be carefully removed of O2 by saturating with high-quality argon. Required materials for the preparation of this solution are as follows: 1. A tank of •NO gas (AirLiquid). 2. Saturated solution of •NO is prepared in deoxygenated ultrapure Milli-Q water containing 100 μM DTPA (diethylenetriaminepentaacetic acid, Sigma) in a glass tube sealed with a Teflon cap (e.g., Vacutainer tube).

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3. The •NO gas is washed from higher oxides by two sequential passages through 10 M NaOH prior to being bubbled in solution vial. 4. Determination of •NO concentration in the stock solution is performed on the ISO-NOP 2 mm Pt sensor connected to an ISO-NO Mark II (World Precision Instruments, Inc., USA). 5. The ISO-NOP sensor is calibrated in accordance to the specifications and required the following solutions: 100 μM NaNO2, 0.2 M KI, and 0.2 M H2SO4. 2.2. Microelectrode Fabrication and Evaluation

2.2.1. Microelectrode Fabrication

The fabrication of microelectrodes for the electrochemical measurement of •NO entails the construction, surface modification (for • NO microsensors), evaluation of general recording properties, and calibration of each microelectrode. The required materials for this process are as follows: 1. Carbon fibers. We use different types of carbon fibers: (a) For •NO microsensors, we use 30 μm diameter fibers from Textron Lowell, MA. (b) For O2 microelectrodes, we use 10 μm diameter fibers from Amoco Corp., Greenville, SC. 2. Borosilicate glass capillaries (1.16 mm i.d. × 2.0 o.d.) from Harvard Apparatus Ltd., UK. 3. Acetone for washing carbon fiber and insertion into capillary. 4. Copper wire (we use individualized copper wires from network cables). 5. Silver conductive paint (RS, UK). 6. Cyanoacrylate glue. 7. Vertical puller (Harvard Apparatus Ltd, UK). 8. A pair of small tweezers (0.5 mm tip). 9. Microscope (inverted will work best).

2.2.2. Modification of Carbon Fiber Microelectrodes

1. Nafion® solution, by Aldrich (5 wt% in a mixture of aliphatic alcohols). 2. PBS lite (10 mM NaH2PO4, 40 mM Na2HPO4, and 100 mM NaCl, pH 7.4). 3. A 5 mM o-PD prepared in deoxygenated PBS lite containing 100 μM ascorbic acid. 4. An oven (must reach 170°C).

2.2.3. Microelectrode Evaluation and Calibration

1. The evaluation and calibration of microelectrodes is performed in PBS lite. 2. The microelectrodes are systematically tested for their general recording properties by Fast Cyclic Voltammetry (FCV) using a potentiostat (Ensman Instruments, USA).

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3. The calibration of O2 microelectrodes requires argon and Carbox (95% O2/5% CO2) to vary (O2) in PBS lite. 4. Calibration is performed in the slice recording chamber in a two electrode mode circuit with an Ag/AgCl pellet as a reference electrode and the microelectrode as the working electrode. Measurement of O2 is performed with a picoamperometer (AMU 130 model, Tacussel, France) connected to a Zipp&Zonen charter. 5. For calibration of •NO microsensor, a saturated •NO stock solution is used. During the calibration of the •NO microsensor, selectivity is also evaluated. Prepare stock solutions of AA and NO2− (20 mM). 6. Calibration is also performed in a two electrode circuit using an Ag/AgCl 3 M reference electrode and the microsensor as the working electrode, in a 40-mL beaker. The electrochemical recordings of •NO are performed within the NO Model T electrochemical detection system (Innovative Instruments, Tampa, Pl, Florida). 2.3. Simultaneous Recording of •NO and O2 in Rat Hippocampal Slices 2.3.1. Preparation of Rat Hippocampal Slices

1. Isolation and recovery of rat hippocampal slices is performed using aCSF (124 mM NaCl, 2 mM KCl, 25 mM NaHCO3, 1.25 mM KH2PO4, 1.5 mM CaCl2, 1.4 mM MgCl2, and 10 mM D-glucose). The aCSF solution must be saturated with Carbox to assure pH buffering and oxygenation (see Note 1). 2. Dissection materials required for preparation of hippocampal slices are as follows: Two medium (10 cm) and one small petri dish (5 cm), one scalpel, two pairs of scissors (one of which should be able to cut scalp bone – if larger animals are used, using serrated scissors may be considered useful), one pair of large forceps (these will be used to hold the decapitated rat head, so the distance of the tips should be close to the distance between animal eyes), two small forceps with curved and serrated tips (tip curvature should be between 45° and 90°; tip diameter is not critical, but around 0.8 mm will be adequate), one rongeur and one spatula, paper filter (to fit the medium petri dish) and clear acetate sheet cut to the size of the tissue chopper stage. 3. Other materials needed are a tissue chopper (McIlwain Tissue Chopper, Campden Instruments, UK) and a tissue slice preincubation chamber (BSC-PC model, Harvard Apparatus). In order to transfer slices from one place to another, a 1- or 5-mL automatic pipette can be used with a disposable tip in which the extremity has been cut to allow aspiration of slices without touching them (approximately 5 mm in diameter). 4. Slice separation: separating slices after sectioning is a critical step in the procedure, as much care should be taken to avoid

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lesion of the tissue. To perform this separation, we use Pasteur pipettes in which the tip has been blunted and slightly curved by heating. Also or alternatively, one can use a very small paintbrush, with only a couple of threads. 2.3.2. Electrochemical Recording of,•NO and O2 in Hippocampal Slices

1. Thermostatic bath is placed above the level of recording chamber to allow solutions to perfuse in a siphon system. 2. Electrochemical recordings in slices are performed in a perfusion recording chamber (BSC-BU with BSC-ZT top, from Harvard Apparatus, USA) coupled to a temperature controller (model TC-202A, Harvard Apparatus, USA). 3. The amperometric recording currents of •NO are performed within the NO Model T system. 4. Concerning the electrochemical recording of O2, amperometric currents are recorded using the AMU 130 picoamperometer connected to a Zipp&Zonen charter. 5. To position the microsensors, a micromanipulator is required. 6. All works done using this recording system require the use of a binocular stereomicroscope on a boom stand. In our setup, we use a SZ60 from Olympus.

3. Methods 3.1. Preparation and Determination of Saturated,•NO Stock Solution

This solution is prepared freshly on the day of use. 1. Remove O2 from the Milli-Q water (containing DTPA) by bubbling argon or N2 for 40 min. 2. To obtain a saturated stock solution, bubble •NO gas for 40 min. After this period, the concentration of •NO is approximately 2.0 mM at room temperature (32). 3. The concentration of this saturated •NO solution is determined by means of the ISO-NOP 2 mm Pt electrode. This is a platinum electrode covered with a gas permeable membrane that confers selectivity of the sensor to •NO. This sensor is calibrated daily using the recommended calibration procedure, by chemical generation of •NO from the following reaction: 2NaNO 2 + 2KI + 2H 2SO 4 ® 2• NO + I 2 + H 2 O + 2Na 2SO 4 Add NaNO2 aliquots to a mixture of 0.1 M KI and 0.1 M H2SO4. In the presence of excess reductant, all NO2− is converted to •NO. 4. After calibrating the ISO-NOP electrode, determine the •NO concentration in deoxygenated PBS lite. Allow current to stabilize

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and then add an aliquot of •NO solution. Repeat this step three times and extrapolate the •NO concentration of the stock solution. 3.2. Carbon Fiber Microelectrodes 3.2.1. Fabrication of Microelectrodes

1. Place capillaries in a petri dish filled with acetone with one extremity of the capillary immersed in the liquid and the other over the opposite edge of the petri dish protruding out of the dish. The capillaries should be filled with acetone. By using a pair of tweezers, insert one individual fiber into each capillary, making sure that the fiber has crossed the whole length. This procedure should be performed near a source of natural light or using a cold light source – convection from lights will make the fibers move in the air, making insertion into the capillary more difficult. 2. Allow the acetone to evaporate (see Note 2). 3. Mount the capillary with a fiber onto the vertical puller. After pulling, retain the half containing the fiber and cut the exposed section to a length of approximately 1 cm from end of glass (see Note 3). 4. Mount the microelectrode on a microscope and cut exposed carbon fiber at desired length using small tweezers. For •NO microsensors, we cut fibers at 100–150 μm of length and for O2, fibers are cut at 10–20 μm (see Note 4). 5. Use a syringe with a long/thin tip to deposit a small amount of conductive silver paint inside the capillary, at the stem end. 6. Insert a thin copper wire in the capillary. Be sure that the copper is exposed at both extremities (remove insulating plastic from tips). The exposed tip must reach the conductive silver paint. 7. To finish the construction process, add a small drop of cyanoacrylate glue at the top end of the capillary to fix the copper wire in position. 8. The general recording properties of the microelectrodes is evaluated in PBS lite by FCV performed between −0.4 and +1.6 V vs. Ag/AgCl at a scan rate of 200 V/s. This potential range offers an electrical pretreatment of the carbon fiber, enhancing sensitivity. The observation of stable background current and sharp transients at the reversal potentials indicates suitable recording properties.

3.2.2. •NO Microsensors

While the electrochemical measurement of O2 can be performed using bare carbon fiber microelectrodes, for •NO detection, the active carbon surface must be modified to achieve sensor selectivity. One of the problematic issues associated with the electrochemical recording of •NO is the high potential required (+0.9 V vs. Ag/ AgCl) to drive oxidation at the electrode surface. At this working potential, many other electroactive species present in biological

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preparations (namely, from nervous system, biogenic amines and indoles, ascorbic acid, and NO2−) can become interferents. One strategy used to increase selectivity is the chemical modification of the active carbon surface of the electrode. Here, the active surface of the carbon fiber is modified by coating with Nafion® and electropolymerization of o-PD. Nafion® is a sulfonated tetrafluoroethylene based fluoropolymer-copolymer acting as an anionic repellant, thus preventing access of molecules such as ascorbic acid to the active surface of the microelectrode. The polymerized o-PD creates an exclusion layer at the carbon fiber surface, minimizing the access of interfering molecules such as ascorbic acid, dopamine, and 5-hydroxytryptamine to the carbon surface and also preventing fouling of the carbon surface (33). 1. The microelectrodes are dried in an oven at 170°C for 4 min to remove traces of humidity. 2. The microelectrode tip is dipped into the Nafion® solution at room temperature for 1–2 s and then dried at 170°C for 4 min. The best results are obtained when two layers of Nafion® are applied (see Note 5). After coating with Nafion®, the microelectrodes can be stored at 4°C for 1 week until usage. 3. The microelectrode coated with Nafion® is placed in a fresh o-PD solution and electropolymerization is achieved by applying a constant potential +0.7 V vs. Ag/AgCl for 30 min, with stirring. The microsensor (coated microelectrode) is then rinsed with deionized water and placed in a 50-mL beaker containing PBS lite. The electrodes are coated and used on the day of use and not reused (see Note 6). Figure 1 shows a schematic representation of a carbon fiber microelectrode and surface modification details. 3.2.3. Calibration of •NO Microsensor

1. Calibration of the •NO microsensor is performed in a 40-mL beaker containing 20 mL deoxygenated PBS lite with gentle stirring. 2. Place •NO microsensor in PBS lite (using a micromanipulator) and polarize the electrode at +0.9 V vs. Ag/AgCl. 3. Allow the oxidation current to stabilize. 4. Add •NO from stock solution. We usually perform three additions of 0.5–1 μM,•NO. 5. In order to evaluate microsensor selectivity, we add 250 μM of ascorbic acid prior to addition of •NO and 100 μM NO2− after addition of •NO. Testing of other possible electroactive interferents is recommended. A typical calibration recording and calibration curve are presented in Fig. 2a.

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Fig. 1. Schematic representation of a carbon fiber microelectrode for •NO (•NO microsensor) showing tip detail, seal between the carbon fiber and the pulled glass capillary as well as chemical modification of carbon surface with Nafion® and o-PD, which increase electrode selectivity toward negatively charged species and large compounds, respectively. AA ascorbic acid; DA dopamine; DOPAC 3,4-dihydroxyphenylacetic acid; 5-HT 5-hydroxytryptamine.

3.2.4. Calibration of O2 Microelectrode

1. The O2 microelectrode is calibrated in the perfusion chamber used for slice recordings. This approach allows the calibrations to be performed at 32°C, the same temperature used for slice recordings. 2. The calibration of the O2 microelectrode requires three solutions at different O2 concentrations. We use PBS lite saturated with argon (0 mM O2), at atmospheric O2 concentration (0.24 mM O2 at 32°C) and saturated with Carbox (1 mM O2 at 32°C). These solutions are preheated in the bath at 34°C and perfused in the recording chamber, at 32°C. The first solution used is the PBS lite at atmospheric (O2). 3. Using a micromanipulator, the microelectrode tip is immersed in the perfusion media in the recording chamber. 4. The microelectrode is polarized at −0.8 V vs. Ag/AgCl pellet and the current is allowed to stabilize. 5. Once a stable current is obtained, switch the perfusing solution to PBS lite 0 mM O2 and finally at 1 mM O2. A typical calibration curve is shown in Fig. 2b.

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Fig. 2. (a) Typical amperometric recording and calibration curve (inset ) of the,•NO microsensor. The microelectrode selectivity is tested by adding ascorbic acid (AA) prior to •NO and NO2− afterward. As shown, the microsensor response to these two intereferents is negligible. (b) Typical amperometric recording and calibrations curve (inset ) of a O2 microelectrode.

3.3. Monitoring of the Interacting Profiles of Endogenous •NO and O2 in Rat Hippocampal Slices 3.3.1. Preparation of Rat Hippocampal Slices

We typically use male Wistar rats with age between 5 and 6 weeks (approximately 150 g). However, the procedure is the same for animals of other ages. 1. Prepare ice-cold aCSF (2 × 50 mL) and fill the preincubation chamber with aCSF at room temperature. Allow both solutions to saturate with Carbox for 40 min. 2. Sacrifice the animal by cervical displacement and decapitate. 3. Use a pair of scissors to cut the skin and expose the cranium. Then, cut through the cranial bones along the sagittal suture and remove the parietal skull bones to each side, exposing the brain. If older animals are used, serrated scissors may be required because cranial bone is very hard. The frontal bone can be

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removed with the assistance of the rongeur. Make sure that the meningeal membranes are removed. 4. With a spatula cut the optical nerves (with one caudal-frontal movement of the spatula inserted under the brain) and then remove the whole brain into a medium petri dish with filter paper. Add ice-cold aCSF saturated with Carbox. One can additionally bubble Carbox in the medium during dissection (see Note 7). 5. With the help of a scalpel, separate the cerebellum from the encephalon and cut the two hemispheres apart. 6. Flip one hemisphere for the cortex to be facing down and the ventral side facing up. With one pair of curved forceps, gently hold the encephalon at the frontal end; using the curved section of the second pair of curved forceps, remove the midbrain and expose the hippocampus. Gently flip the hippocampus over and cut along the subiculum. Repeat this process with the other hemisphere. 7. Place the hippocampi on the stage of the tissue chopper (on the acetate film) with the long axis perpendicular to the blade. The tissue should be dry and the blade slightly wet. 8. Cut 400-μm sections (see Note 8). 9. Transfer sectioned hippocampi into a small petri dish (transfer the acetate with the tissue into the dish) and add ice-cold aCSF saturated with Carbox. 10. Separate the slices using Pasteur pipettes with sealed end or single thread paintbrush. This is a critical step, and care should be taken not to damage the tissue. 11. Using the 1- or 5-mL cut pipette tip, transfer separated slices into the prerecording chamber containing room temperature aCSF continuously bubbled with Carbox. Discard slices that are not intact (during dissection and separation, the inferior blade of the dentate gyrus may be lost or cuts to the hippocampus may disrupt the tissue integrity). 12. Slices are maintained in the prerecording chamber for at least 1 h prior to use, allowing for good tissue recovery. 3.3.2. Simultaneous Recording of •NO and O2 Concentration Dynamics in Rat Hippocampal Slices

1. Solutions perfused in the recording chamber are preheated at 34°C to prevent formation of gas bubbles in the tubing and chamber, which cause interference in electrochemical recordings. Before placing a slice in the recording chamber, make sure that the temperature has stabilized – temperature fluctuations alter recorded oxidation and reduction currents. 2. Transfer one hippocampal slice into the recording chamber and fix it to the nylon mesh (see Note 9).

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3. Place each microelectrode on the holder of a micromanipulator and insert them into the CA1 subregion of the hippocampal slice (or other region of interest). The microelectrodes are inserted at a 45°–60° angle and the exposed tip is placed at a depth of 200–300 μm into the tissue (see Note 10). Figure 3 shows a schematic representation of the setup used to perform these simultaneous recordings. 4. For high temporal and spatial resolution of the simultaneous recording, the distance between the electrodes should be as small as possible (<50 μm). 5. Allow both current recordings to stabilize and perform tissue stimulation. Typically, NMDA is perfused at concentrations 10, 50, and 100 μM during 2 min. NMDA is added to carbox saturated aCSF in a different beaker and the perfusion solution is switched using a stopcock. This stimulation protocol results in a transient increase in •NO concentration in the hippocampal slice (in particular in the CA1 subregion) and allows for the

Fig. 3. Schematic representation of the setup used in the simultaneous recording of •NO and O2 concentration dynamics in the hippocampal slice. In (a), a lateral view of the recording chamber top is shown, with the electrodes placed in the tissue and instrumentation used. In (b), a top view showing the two electrodes placed in the CA1 subregion of the hippocampal slice.

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study of how different •NO concentration dynamics affect O2 consumption in the tissue. Figure 4 shows typical recording of • NO and O2 in the CA1 subregion of the hippocampal slice stimulated with 10, 50, and 100 μM NMDA. 6. Pharmacological modulation can be achieved either by pretreating slices or by addition to perfusion media (see Note 11). 3.4. Concluding Remarks

The approach here described allows for the study of •NO regulation of cellular respiration in a tissue model closer to the in vivo setting, thus offering several advantages over simpler systems such as mitochondrial suspensions and cell cultures. The simultaneous, direct, and real-time recording of •NO and O2 concentration

Fig. 4. Typical recording of •NO and O2 in the CA1 subregion of the hippocamal slice challenged with NMDA 100 μM (a), 50 μM (b), and 10 μM (c) applied for a 2-min period in perfusion (box). The solid line is the •NO oxidation current and dashed line is the O2 reduction current. Stimulation of the tissue with NMDA induces concentration-dependent increase in •NO. The evolution of the O2 profile is more complex: activation of the neuronal tissue induces an initial increase in O2 consumption (1) which is followed by a decrease in consumption (2) until a new steady state is reached (3). The analysis of these and other simultaneous recordings (for more information, see ref. 16) revealed that the maximal variation in (O2) was directly related to the peak (•NO) and that a threshold concentration of •NO was required for inhibition of O2 consumption to be observed. Together, the results indicated that NMDA-evoked •NO production inhibitied O2 consumption in the CA1 subregion of the hippocampal slice in a concentration-dependent fashion (reproduced from ref. 16 with permission from Elsevier).

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profiles in a biological preparation that retains high cytoarchitectural integrity is of great interest in gaining deeper insight of the mechanism by which endogenous •NO regulates mitochondrial respiration in vivo. Among other reasons, both gases diffuse into the tissue (similar to in vivo) and O2 tension is physiological, turning the direct reaction between the two gases meaningless. The approach described here may be applied not only for the study of mitochondrial metabolism but also in the context of mitochondrial pharmacological studies. Understanding how different • NO concentration dynamics impacts mitochondrial respiration, be it the reversible inhibition of CcO by •NO, the irreversible inhibition of other mitochondrial complexes by peroxynitrite, or even mitochondrial production of reactive oxygen species with signaling properties (such as H2O2 and O2−•), not only is interesting from the standpoint of fundamental research but may also be useful in pharmacological studies and even in disease models where mitochondrial dysfunction is proposed to occur.

4. Notes 1. For the isolation of rat hippocampal slices, ice chips can be allowed to form by placing aCFS at −20°C for a short period. The low temperature is essential to guarantee higher viability of tissue during isolation process. 2. The acetone serves two purposes: first, it facilitates the insertion of the fiber into the capillary (this is essential when using the 10 μm fiber, otherwise it will not slide into the capillary); second, it cleans the carbon fiber of residues. 3. Pulling of the capillary/fiber array is a critical step in the fabrication of carbon fiber microelectrodes. The strength and heat of the pulling must be adjusted to obtain a tight seal between the capillary and the carbon fiber. Also, the pulled portion of the glass should not be excessively long to avoid bending of the microelectrode tip. 4. The carbon fibers are cut at different lengths in accordance to the purpose of use. In the case of •NO microsensors, the fiber is cut longer to increase recording surface and thus sensitivity. For O2 microelectrodes, the fiber is cut short to increase spatial resolution of the recording. 5. Lower temperature curing produces thicker films compared to higher temperature curing (34), so it is important do guarantee that Nafion® is dried at 170°C. The application of Nafion®, while increasing electrode selectivity, will also decrease sensitivity and response time. As such, a compromise must be reached between these properties. Others and we have established

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that the application of two layers of Nafion® results in good selectivity without significantly compromising sensitivity and response time (27, 33, 35). 6. After the Nafion® film has been hydrated, it must not be allowed to dry. Thus, after polymerization of o-PD, the electrode must always remain with the tip immersed in PBS. 7. It is critical to reduce to a minimum the time between the death of the animal and having the brain submerged in ice-cold aCSF. Ideally, this procedure should last no longer than 1 min. During this period, the nervous tissue is not being oxygenated and elevated temperature leads to high metabolic activity. 8. Using sections at 400 μm of thickness guarantees that the core of the tissue is healthy and sufficiently oxygenated during recordings at 32°C. During the sectioning process, about 100 μm of tissue on each side of the slice is damaged (30, 31). 9. Holding the slices in the nylon mesh may be a difficult task. It is useful to gently scratch the nylon mesh with a needle. To fix the slice, the fimbria can be crushed against the mesh. 10. It is important to perform recordings in the tissue core, as the upper and lower 100 μm of tissue are damaged due to slice preparation. Also, at the tissue core, the basal (O2) is at physiological levels (29, 31, 36). 11. Any compound applied to the perfusion media must be tested for interference at the microelectrodes.

Acknowledgment This work was funded by grant PTDC/SAU-NEU/108992/2008 from FCT (Portugal). References 1. Moncada S, Higgs A (1993) The L-arginine-nitric oxide pathway. N Engl J Med 329:2002–2012 2. Poulos TL (2006) Soluble guanylate cyclase. Curr Opin Struct Biol 16:736–743 3. Kleppisch T, Feil R (2009) cGMP signalling in the mammalian brain: role in synaptic plasticity and behaviour. Handb Exp Pharmacol 191:549–579 4. Moncada S, Higgs EA (2006) Nitric oxide and the vascular endothelium. Handb Exp Pharmacol 176(1):213–254 5. Bolanos JP, Peuchen S, Heales SJ, Land JM, Clark JB (1994) Nitric oxide-mediated inhibition of the mitochondrial respiratory

chain in cultured astrocytes. J Neurochem 63:910–916 6. Brown GC, Cooper CE (1994) Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett 356:295–298 7. Cleeter MW, Cooper JM, Darley-Usmar VM, Moncada S, Schapira AH (1994) Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. Implications for neurodegenerative diseases. FEBS Lett 345:50–54

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8. Schweizer M, Richter C (1994) Nitric oxide potently and reversibly deenergizes mitochondria at low oxygen tension. Biochem Biophys Res Commun 204:169–175 9. Brudvig GW, Stevens TH, Chan SI (1980) Reactions of nitric oxide with cytochrome c oxidase. Biochemistry 19:5275–5285 10. Gibson QH, Greenwood C (1963) Reactions of cytochrome oxidase with oxygen and carbon monoxide. Biochem J 86:541–554 11. Stevens TH, Brudvig GW, Bocian DF, Chan SI (1979) Structure of cytochrome a3-Cua3 couple in cytochrome c oxidase as revealed by nitric oxide binding studies. Proc Natl Acad Sci USA 76(7):3320–3324 12. Wainio WW (1955) Reactions of cytochrome oxidase. The Journal of biological chemistry 212(2):723–733 13. Antunes F, Boveris A, Cadenas E (2004) On the mechanism and biology of cytochrome oxidase inhibition by nitric oxide. Proc Natl Acad Sci USA 101:16774–16779 14. Antunes F, Cadenas E (2007) The mechanism of cytochrome C oxidase inhibition by nitric oxide. Front Biosci 12:975–985 15. Cooper CE, Mason MG, Nicholls P (2008) A dynamic model of nitric oxide inhibition of mitochondrial cytochrome c oxidase. Biochimica et biophysica acta 1777:867–876 16. Ledo A, Barbosa R, Cadenas E, Laranjinha J (2010) Dynamic and interacting profiles of *NO and O2 in rat hippocampal slices. Free Radic Biol Med 48:1044–1050 17. Jarrard LE (1995) What does the hippocampus really do? Behav Brain Res 71(1–2):1–10 18. Garthwaite J, Garthwaite G, Palmer RM, Moncada S (1989) NMDA receptor activation induces nitric oxide synthesis from arginine in rat brain slices. Eur J Pharmacol 172(4–5):413–416 19. Prast H, Philippu A (2001) Nitric oxide as modulator of neuronal function. Prog Neurobiol 64(1):51–68 20. Hopper RA, Garthwaite J (2006) Tonic and phasic nitric oxide signals in hippocampal long-term potentiation. J Neurosci 26:11513–11521 21. Laranjinha J, Ledo A (2007) Coordination of physiologic and toxic pathways in hippocampus by nitric oxide and mitochondria. Front Biosci 12:1094–1106 22. Zhuo M, Hawkins RD (1995) Long-term depression: a learning-related type of synaptic plasticity in the mammalian central nervous system. Rev Neurosci 6:259–277 23. Zorumski CF, Izumi Y (1998) Modulation of LTP induction by NMDA receptor activation and nitric oxide release. Prog Brain Res 118: 173–182

24. Amaral DG, Witter MP (1989) The three-dimensional organization of the hippocampal formation: a review of anatomical data. Neuroscience 31:571–591 25. Anderson P, Bliss TV, Skrede KK (1971) Lamellar organization of hippocampal pathways. Exp Brain Res 13:222–238 26. Bliss TV, Collingridge GL (1993) A synaptic model of memory: long-term potentiation in the hippocampus. Nature 361:31–39 27. Ferreira NR, Ledo A, Frade JG, Gerhardt GA, Laranjinha J, Barbosa RM (2005) Electrochemical measurement of endogenously produced nitric oxide in brain slices using Nafion/o-phenylenediamine modified carbon fiber microelectrodes. Analytica Chimica Acta 535:1–7 28. Gerhardt GA, Oke AF, Nagy G, Moghaddam B, Adams RN (1984) Nafion-coated electrodes with high selectivity for CNS electrochemistry. Brain Res 290:390–395 29. Ledo A, Barbosa RM, Gerhardt GA, Cadenas E, Laranjinha J (2005) Concentration dynamics of nitric oxide in rat hippocampal subregions evoked by stimulation of the NMDA glutamate receptor. Proc Natl Acad Sci USA 102: 17483–17488 30. Jiang C, Agulian S, Haddad GG (1991) O2 tension in adult and neonatal brain slices under several experimental conditions. Brain Res 568:159–164 31. Mulkey DK, Henderson RA 3rd, Olson JE, Putnam RW, Dean JB (2001) Oxygen measurements in brain stem slices exposed to normobaric hyperoxia and hyperbaric oxygen. J Appl Physiol 90:1887–1899 32. Zacharia IG, Deen WM (2005) Diffusivity and solubility of nitric oxide in water and saline. Ann Biomed Eng 33:214–222 33. Friedemann MN, Robinson SW, Gerhardt GA (1996) o-Phenylenediamine-modified carbon fiber electrodes for the detection of nitric oxide. Anal Chem 68:2621–2628 34. Gerhardt GA, Hoffman AF (2001) Effects of recording media composition on the responses of Nafion-coated carbon fiber microelectrodes measured using high-speed chronoamperometry. J Neurosci Methods 109:13–21 35. Santos RM, Lourenco CF, Piedade AP, Andrews R, Pomerleau F, Huettl P, Gerhardt GA, Laranjinha J, Barbosa RM (2008) A comparative study of carbon fiber-based microelectrodes for the measurement of nitric oxide in brain tissue. Biosens Bioelectron 24:704–709 36. Erecinska M, Silver IA (2001) Tissue oxygen tension and brain sensitivity to hypoxia. Respir Physiol 128:263–276

Chapter 6 Mitochondrial Membrane Potential (DY) Fluctuations Associated with the Metabolic States of Mitochondria Carlos M. Palmeira and Anabela P. Rolo Abstract The study of the mitochondrial membrane potential (ΔΨ) is essential for an integrated appraisal of mitochondrial function, since it reflects differences in electrical potential and represents the main component of the proton electrochemical gradient accounting for more than 90% of the total available respiratory energy. Numerous methods have been used to estimate mitochondrial membrane potential (ΔΨ), including fluorescent methods and electrochemical probes. In this chapter, we describe several practical approaches that allow mitochondrial membrane potential (ΔΨ) evaluation, by using a tetraphenylphosphonium (TPP+)-selective electrode. The main focus is given to the evaluation of ΔΨ in isolated mitochondria. Key words: TPP+-selective electrode, Membrane potential, Mitochondria, Permeabilized cells, Metabolic states, Permeability transition

1. Introduction The inner mitochondrial membrane transduces energy through oxidative phosphorylation. This is the main process responsible for the production in the form of ATP in eucaryotic cells (1). The entire general sequence of biochemical events leading to ATP synthesis has been known since Mitchell proposed his chemiosmotic theory (2). It elegantly described how mitochondrial respiration creates an electrochemical gradient of protons across the mitochondrial inner membrane, which in turn drives ATP synthesis through the mitochondrial ATP synthase. However, the way in which the oxidative phosphorylation system is regulated in intact tissues still remains a matter of debate. Besides ATP production, other mitochondrial activities that require energy, such as electrophoretic

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_6, © Springer Science+Business Media, LLC 2012

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or protonophoric transport of ions, metabolic substrates, and proteins for the mitochondrial matrix, are supported by the primary form of energy generated in mitochondria, the electrochemical proton gradient. Interference with the generation of the electrochemical proton gradient or its induced dissipation affects mitochondrial bioenergetics. The initial event of energy conservation is charge separation at the inner mitochondrial membrane. The electrochemical proton gradient is generated by means of electrogenic pumping of protons, from the mitochondrial matrix to the intermembrane space, which is catalyzed by the respiratory chain complexes (Fig. 1). Electrons deriving from oxidation of substrates are funneled through the redox carriers of the respiratory chain (3). This process is coupled to proton ejection at complexes I (NADH:ubiquinone reductase), III (ubiquinol: cytochrome c reductase or bc1-complex) and IV (cytochrome c oxidase). The final electron acceptor is molecular oxygen, which through four electron reduction is converted to water. The succession of electron transfer occurs in the following sequence: complex I – ubiquinone – complex III – cytochrome c – complex IV – O2. The amplitude of the electrochemical proton gradient, which is known as respiratory control, regulates the overall rate of electron transport in the respiratory chain. Ubiquinone is a mobile electron carrier, dissolved in the lipid phase of the membrane, and interacts specifically with complexes I and III (4). Succinate dehydrogenase receives electrons from succinate (of the tricarboxylic acid cycle), through the oxidation of FADH2, reducing ubiquinone to ubiquinol. Cytochrome c,

Fig. 1. Mitochondrial electron transport chain and dissipation of the proton gradient by UCPs. CI complex I, CII complex II, CIII complex III, CIV complex IV, CV (complex V) or ATPsynthase. TCA tricarboxylic acid cycle, UCP uncoupling protein.

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a mobile protein attached to the cytosolic face of the inner mitochondrial membrane, serves as an electron carrier between complexes III and IV. Energy released from the oxidation of substrates in the matrix is used to reduce NAD+ and ubiquinone, originating NADH and ubiquinol. NADH is oxidized by complex I, which is composed of more than forty polypeptides, seven of which are encoded in the mitochondrial genome. Complex I contains a prosthetic flavin mononucleotide and six Fe-S centers. It also contains a binding site for ubiquinone that receives reducing equivalents in steps of one electron forming transient semiquinone radicals. Rotenone is a lipophilic pesticide that binds with high affinity to complex I, specifically inhibiting its catalytic activity, that is, it inhibits the transfer of electrons from complex I to ubiquinone (5). Since this is the main entry point of the respiratory chain, inhibition of complex I block most of the oxidative metabolic reactions conducted by mitochondria. Complex III oxidizes the reduced ubiquinol, being the second entry point of the respiratory chain. Antimycin A inhibits the transfer of electrons from complex II to cytochrome c. Cytochrome b, one Fe-S protein, and an hydrophobic cytochrome c1 are the main polypeptides that anchor the redox centers. Complex IV (three subunits encoded by mitochondrial DNA) is inhibited by cyanide; NO• inhibits complex IV in a reversible way, through competition with O2. Complex IV uses more than 90% of the oxygen taken up by the cell. The terminal reduction of O2 is processed in two steps: (1) transient formations of oxide anions ( O 2 - ) in the active site of the enzyme and (2) the reaction of O 2 - with matrix protons, promoting the formation of H2O. This two-step reaction avoids the formation of superoxide radical anion ( O•2- ), since it does not occur release of partially reduced oxygen species because of the high binding affinity of cytochrome c oxidase. However, during normal metabolism, about 1–5% of the 90% of oxygen is converted into superoxide. The production of toxic reactive oxygen species is significantly increased with inhibition of complex III or IV. The reduction of O2 to H2O also induces matrix alkalinization, which helps establishing the transmembrane electrochemical proton gradient. The electrochemical proton gradient, which forms the so-called protonmotive force (Δp), consists of the electrical membrane potential (ΔΨ) and the pH gradient (ΔpH), across the inner mitochondrial membrane. The magnitude of the electrochemical proton gradient is about −220 mV, and under physiological conditions most of the gradient is in the form of the ΔΨ (6). Because the matrix side of the inner mitochondrial membrane is negatively charged and slightly alkaline, mitochondria can accumulate large amounts of positively charged lipophilic compounds and some acids. Complex V or ATP synthase uses the electrochemical proton gradient as the driving force to synthesize ATP from ADP and phosphate (2, 7, 8). Complex V can also operate in reverse, as a proton-translocating ATPase. The ATP

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Fig. 2. Schematic representation of TPP + distribution across the mitochondrial membrane, according with the Nernst equation.

synthase complex consists of two assemblies, with a variety of polypeptide subunits (only two encoded by mitochondrial DNA). The extrinsic F1 contains the catalytic sites. The membrane assembly Fo is a proton channel, specifically inhibited by oligomycin. The return of protons through Fo, activates the catalytic sites in F1 to phosphorylate ADP. The rate of mitochondrial ATP synthesis is regulated by alterations in the ATP/ADP ratio (phosphorylation potential) and the NADH/NAD+ ratio (redox potential) (9). 1.1. Estimation of DY

Since TPP+ equilibrates passively across biological membranes until electrochemical equilibrium is reached DΨ can be estimated from the Nernst equation: DY (mV) = 59 log [TPP + ] in / [TPP + ] out , at 25°C where [TPP+]in and [TPP+]out represent the concentrations of the ion in the matrix phase and in the medium, respectively (Fig. 2). Possible differences in activity coefficients are neglected. The internal volume of mitochondria is taken as 1.1 μl/mg protein. Alternatively, the membrane potential can be estimated from the following equation (at 25°C): DY (mV ) = 59 log (v / V ) - 59 log (10DE / 59 - 1)

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where v, V, and ΔE are mitochondrial volume, volume of the incubation medium, and deflection of the electrode potential from the baseline. Changes in the membrane potential when mitochondria go from state a to state b are given by: DY /59 = log (10 DEa /59 - 1) - log (10DEb/59 - 1) which shows that it is necessary to measure ΔEa and ΔEb and not just the difference Ea − Eb. TPP+ must be included in the reaction solution at a concentration of 3 μM to achieve high sensitivity in measurements and to avoid possible toxic effects on mitochondria (10).

2. Materials 2.1. Reagents and Buffers

Tetraphenylphosphonium, tetraphenylboron, diisooctylphthalate, substrates of respiratory chain (glutamate/malate and succinate), inhibitors of respiratory chain (rotenone, cyanide), ionophores (Nigericin, FCCP), inhibitors of TCA cycle (salicylate, Br-succinimide), inhibitors of the adenine nucleotide translocator (atractyloside), inhibitor of the calcium uniporter (ruthenium red) are all from Sigma (St. Louis, MO). 1. Reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4.

2.2. Isolation of Mitochondrial Fraction Solutions

1. Medium A (homogenization medium): 225 mM mannitol, 75 mM sucrose, 0.5 mM EGTA, 0.5 mM EDTA, 0.1% BSA (fatty acid free), 10 mM HEPES, pH 7.4. 2. Medium B (washing medium): 225 mM mannitol, 75 mM sucrose, 10 mM HEPES, pH 7.2.

2.3. Preparation of TPP+ Electrode

Membrane potentials are measured with a TPP+ electrode constructed by the use of a polyvinylchloride-based membrane containing tetraphenylboron as an ion-exchanger, prepared according to Kamo et al., using a calomel electrode as the ref. 11. The polyvinylchloride membrane is prepared by allowing a solution containing 0.34 mg of tetraphenylboron (Na+ salt), 16 mg of polyvinylchloride (high molecular weight), 57 μl of diisooctylphthalate, and tetrahydrofuran (to a final volume of 500 μl) to evaporate on a glass plate constrained by a glass ring of 1.9-cm diameter. The ring must be covered with a glass beaker and tetrahydrofuran evaporate overnight at room temperature. A clear membrane (0.2-mm thick) is obtained.

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A piece of membrane is glue with tetrahydrofuran to a polyvinyl chloride tube having an inner diameter of about 2 mm. Care must be taken to avoid that tetrahydrofuran causes extensive dissolution of the central part of the membrane through which the TPP+ concentration is sensed. Light sucking and blowing into the tube is carried out till tetrahydrofuran is evaporated. Any membrane material overlapping the tubing is cut out with a razor blade or a special scissor. The complete electrode is filled with 0.1–0.2 ml TPP+ 10 mM as reference solution (the solution should be first degassed) in which a silver wire, coated with AgCl, made the connection with a suitable electrometer. Before use, it is necessary that the electrode is soaked overnight in a 10 mM TPP+ solution for conditioning. The electromotive force is measured between the TPP+ electrode and a calomel electrode in the sample solution. A good electrode must have a voltage response to log [TPP+] linear with a slope of 59 at 25°C, in agreement with the Nernst equation. A rapid observation of the good condition of the electrode can be detected by successive additions of a 1 mM TPP+ solution, doubling each time the preceding concentration. As ΔE = 2.3RT/nF log[C1/C2], if C1/C2 is 2 and 2.3 RT/nF is 59 mV, then ΔE = 17.8 mV, and (for instance) to a recorder deviation of 200 mm for 20 mV, ΔE must be 178 mm for each doubling of the TPP+ concentration.

3. Methods 3.1. Isolation of Rat Liver Mitochondria

1. Take one rat (250 g), starved overnight, and kill it by decapitation. Bleed the rat into the sink for about 1 min (12). 2. Using scissors, cut across the abdomen under the rib cage, remove the liver as soon as possible and drop it into a beaker containing ice-cold homogenization medium (medium A). Remove as much as possible the adhering fat or fibrous tissue, chop up the liver with scissors, and wash the tissue several times in ice-cold homogenizing medium to remove as much blood as possible; the final washing medium should be free of blood. 3. Add approximately 6 ml of cold medium A to each gram of chopped liver and transfer to a precooled glass Potter–Elvejhem homogenizer. 4. Homogenize the tissue using 3–4 up and down strokes of the pestle rotating at 300 rpm (the pestle must reach the bottom of the tube in the first or second pass). 5. Transfer the homogenate to two centrifuge tubes and centrifuge at 800 × g for 10 min in a refrigerated centrifuge to sediment nuclei, red cells, broken and unbroken cells.

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6. Carefully decant the supernatant (leave a small amount of supernatant to avoid contamination of the mitochondrial pellet with nuclei, etc.) and centrifuge it at 10,000 × g for 10 min. 7. Discard the supernatant as completely as possible and gently resuspend the pellet using a test tube containing ice. The mitochondrial pellet form a soft brown pellet; if a red spot is seen in its center, it should be discarded as it consists of pelleted red blood cells. It also generally has a superficial mobile layer (the “fluffy layer”), which must be discarded (it comes off easily upon decanting) together with the supernatant, since it contains damaged and partly broken mitochondria. 8. Recentrifuge the suspension at 10,000 × g for 10 min. 9. Resuspend the pellet in medium B and sediment once more. 10. Suspend carefully the pellet in about 1 ml of medium B, by using a small brush. 3.2. DY Fluctuations Associated to the Phosphorylation– Dephosphorylation Cycle 3.2.1. Effect of FCCP

The membrane potential (ΔΨ) fluctuations (Fig. 3) are measured in an open thermostated vessel, at a constant temperature of 25°C, under efficient magnetic stirring (Fig. 4). 1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min). 3. Add 1 mg mitochondria (this amount can be decreased to 0.5 mg if necessary). Wait 5 min (to allow TPP+ equilibrium). Add 5 mM succinate. Another option is the addition of succinate (or other respiratory substrate of interest) first and start the trace with the addition of 1 mg mitochondria. 4. After TPP+ has been accumulated, add 200 nmol ADP. The potential suddenly decreases upon addition of ADP but, after a short lag phase, the mitochondrial membrane repolarizes close to its state 4 value (before the addition of ADP). Soon after repolarization is completed add KCN 1 mM (inhibitor of cytochrome oxidase complex IV) or FCCP (uncoupler).

3.2.2. Effect of Phosphate

Repeat the experiment above but using medium without phosphate. After the accumulation of TPP+, upon addition of succinate 5 mM, add 5 mM phosphate.

3.2.3. Effect of Nigericin

Repeat the experiment but add 0.5 μg nigericin after the accumulation of TPP+ upon addition of succinate. Nigericin allows an exchange of protons for K+ and the effect means a conversion of ΔpH to ΔΨ. The effect of nigericin is seen as an increased membrane potential.

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Fig. 3. Typical traces obtained with a TPP+ electrode. After the addition of the compound X, we observe a decrease in the initial mitochondrial membrane potential together with an increase in lag phase. This type of trace indicates an effect of this compound at the level of the phosphorylative system.

From the degree of shift upon addition of nigericin, the pH gradient can be estimated. The ΔpH is low, at most 0.2–0.3 pH unit. 3.2.4. Energization of Mitochondrial Membrane by ATP

In this case, choose a chart speed of 1 cm/min and a full scale of 20 mV. Study the energization of mitochondrial membrane upon addition of 2 mM ATP. Soon after TPP+ has been accumulated upon addition of ATP, add 2 μg oligomycin (inhibitor of ATP synthetase). This experiment must be carried out in the presence of rotenone (2 μM) to inhibit the redox chain. 1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone and 3 μM TPP+.

3.3. Examine the Effects of Inhibitors of the TCA Cycle on Membrane Potential

2. Add sequentially 1 mg of protein, 20 μM CaCl2, and 1 mM acetoacetate.

3.3.1. Effect of Salicylate or Br-Succinimide

3. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale (select a chart speed of 2 cm/min). 4. Start the reaction, recording continuously, by adding 1.5 μM of palmitoyl-D,L-carnitine (see the developing of the membrane

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Fig. 4. Image of the TPP+ electrode and experimental setup. The membrane potential ΔΨ) fluctuations are measured in an open thermostated (with water bath at 25°C) incubation chamber with a TPP+ electrode inserted, coupled to a calomel reference electrode and efficient magnetic stirring.

potential). After a steady-state distribution of TPP+ had been reached, add 1.5 mM of salicylate or 1.5 mM of Br-succinimide (inhibitors of TCA cycle). 3.3.2. Effect of Malonyl-CoA

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Add sequentially 1 mg of protein, 20 μM CaCl2, and 1 mM acetoacetate. 3. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale (select a chart speed of 2 cm/min). 4. Start the reaction, recording continuously, by adding 1.5 μM of palmitoyl-CoA and then add 2 mM of L-carnitine. To finalize your assay, add 50 μM malonyl-CoA and record for more 3 min.

3.3.3. Effect of CoASH

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+.

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2. Add sequentially 1 mg of protein, 20 μM CaCl2, and 1 mM acetoacetate. 3. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale (select a chart speed of 2 cm/min). 4. After a steady-state distribution of TPP+ had been reached add 1.5 μM of palmitate + 25 μM CoASH. Add 2 mM of L-carnitine and to finalize the assay, add 1 mM ATP and record for more than 3 min. 3.4. Examine the Effects of Adding Transportable Metabolites to Intact Mitochondria on Membrane Potential

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+.

3.4.1. Effect of Atractyloside

3. Add 1 mg mitochondria (this amount can be decreased to 0.5 mg if necessary). Wait 5 min (to allow TPP+ equilibrium). Add 5 mM succinate. Another option is the addition of succinate (or other respiratory substrate of interest) first and start the trace with the addition of 1 mg mitochondria.

2. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min).

4. After a steady-state distribution of TPP+ had been reached, add 200 nmol ADP. After mitochondrial membrane potential had been reestablished, add 5 μM atractyloside (inhibitor of the adenine nucleotide translocator ANT), followed by 200 nmol ADP and record for an additional 3 min. 3.4.2. Effect of Ruthenium Red

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Place pen recorder on the left edge of the paper recorder after choosing a 50-mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min). 3. Add 1 mg mitochondria (this amount can be decreased to 0.5 mg if necessary). Wait 5 min (to allow TPP+ equilibrium). Add 5 mM succinate. Another option is the addition of succinate (or other respiratory substrate of interest) first and start the trace with the addition of 1 mg mitochondria. 4. After a steady-state distribution of TPP+ had been reached, add 30 μM CaCl2. After mitochondrial membrane potential had been reestablished, add 750 nM Ruthenium Red (inhibitor of the mitochondrial calcium uniporter), followed by 30 μM CaCl2 and record for an additional 3 min.

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1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Place pen recorder on the left edge of the paper recorder after choosing a 50 mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min). 3. Add 1 mg mitochondria (this amount can be decreased to 0.5 mg if necessary). Wait 5 min (to allow TPP+ equilibrium). Add 5 mM succinate. Another option is the addition of succinate (or other respiratory substrate of interest) first and start the trace with the addition of 1 mg mitochondria. 4. After a steady-state distribution of TPP+ had been reached, add 20 μM CaCl2 and 1 mM acetoacetate, followed by 1.5 μM of palmitate + 25 μM CoASH. 5. Add 2 mM of L-carnitine and to finalize the assay, add 0.5 μg/ml oligomycin (inhibitor of ATP synthetase) and record for more 3 min.

3.4.4. Effect of Co-transport

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Place pen recorder on the left edge of the paper recorder after choosing a 50 mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min). 3. Add 1 mg mitochondria (this amount can be decreased to 0.5 mg if necessary). Wait 5 min (to allow TPP+ equilibrium). Add 5 mM succinate. Another option is the addition of succinate (or other respiratory substrate of interest) first and start the trace with the addition of 1 mg mitochondria. 4. Start the reaction, recording continuously, by adding 10 mM glutamate (no development of membrane potential). Add 10 mM Malate (see the developing of the membrane potential) and record for more than 2 min.

3.5. Effects of Oxidants on the Induction of the MPT Pore: Reversal with NEM, DTT (Reducing Agents) or CyA (Inhibitor of the Permeability Transition Pore)

1. Add to the reaction chamber 1 ml of reaction medium containing 130 mM sucrose, 50 mM KCl, 5 mM MgCl2, 5 mM KH2PO4, 10 mM HEPES, pH 7.4, 2 μM rotenone, and 3 μM TPP+. 2. Place pen recorder on the left edge of the paper recorder after choosing a 50 mV full scale; wait until the trace is stable (select a chart speed of 2 cm/min). 3. Add sequentially 5 mM Succinate and 0.5 μg/ml oligomycin. 4. Start the reaction, recording continuously, by adding 1 mg of protein (see the developing of the membrane potential).

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5. After a steady-state distribution of TPP+ had been reached, add 30 μM Ca2+. After mitochondrial membrane potential had been reestablished, add the oxidant (100 μM tert-butyl hydroperoxide or 50 μM Menadione) and record for an additional 5 min. When check for protection with dithiothreitol (DTT; 1 mM), N-ethylmaleimide (NEM; 20 μM), both reducing agents or the inhibitor of the permeability transition pore Cyclosporine A (0.85 μM), add the compounds to the reaction medium at the beginning of the assay (before addition of protein).

4. Notes Lifetime of the electrodes. The parent membranes could be stored dry for years and be used as stock. The average lifetime of the TPP+ selective electrodes is about 2 months. Some general rules for the use of ion-specific electrodes : Generally, to obtain a stable electrode potential and a low background noise care must be taken in the following: 1. Shielding all electrical electrode connections (with aluminum foil). 2. Protecting the reaction vessel with a Faraday cage. 3. Maintaining a constant temperature and stirring. 4. Using a medium with reasonable ionic strength (e.g., 0.1 M KCl). 5. Maintaining a sufficient pH buffering capacity. Hydrophobic substances will cause problems because they might be absorbed by the PVC membrane. This problem may be overcome by the following: 6. Mixing the hydrophobic substances with biological membrane material before bringing it into contact with the electrode. 7. Using these substances in amounts as low as possible (e. g., inhibitors, uncouplers, and organic solvents used to dissolved them in). 8. Washing the electrode after each experiment with ethanol or biological membrane material. Ethanol washing shortens the lifetime of the electrode by extracting some ionophore from the PVC membrane. However, this does not affect the potential, or slope of the electrode, since the membrane contains about a thousand times more ionophore than needed for optimal functioning of the electrode.

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References 1. Saraste M (1999) Oxidative phosphorylation at the fin de siècle. Science 283:1488–1493 2. Mitchell P (1966) Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol Rev 41:445–502 3. Esposti MD, Ghelli A (1994) The mechanism of proton and electron transport in mitochondrial complex I. Biochim Biophys Acta 1187:116–120 4. Trumpower BL (1990) The protonmotive Q cycle. Energy transduction by coupling of proton translocation to electron transfer by the cytochrome bc1 complex. J Biol Chem 265: 11409–11412 5. Esposti MD (1998) Inhibitors of NADHubiquinone reductase: an overview. Biochim Biophys Acta 1364:222–235 6. Azzone GF, Petronilli V, Zoratti M (1984) ‘Cross-talk’ between redox- and ATP-driven H + pumps. Biochem Soc Trans 12:414–416 7. Abrahams JP, Leslie AG, Lutter R, Walker JE (1994) Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370:621–628

8. Capaldi RA, Aggeler R (2002) Mechanism of the F(1)F(0)-type ATP synthase, a biological rotary motor. Trends Biochem Sci 27: 154–160 9. Erecinska M, Wilson DF (1982) Regulation of cellular energy metabolism. J Membr Biol 70:1–14 10. Jensen BD, Gunther TE (1984) The use of tetraphenylphosphonium (TPP+) to measure membrane potentials in mitochondria: membrane binding and respiratory effects. Biophys J 49:105–121 11. Kamo N, Muratsugu R, Hongoh R, Kobatake V (1979) Membrane potential of mitochondria measured with an electrode sensitive to tetraphenyl phosphonium and relationship between proton electrochemical potential and phosphorylation potential in steady state. J Membr Biol 49:105–121 12. Palmeira CM, Moreno AJ, Madeira VMC (1994) Interactions of herbicides 2,4-D and dinoseb with liver mitochondrial bioenergetics. Toxicol Appl Pharmacol 127:50–57

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Chapter 7 Safranine as a Fluorescent Probe for the Evaluation of Mitochondrial Membrane Potential in Isolated Organelles and Permeabilized Cells Tiago R. Figueira, Daniela R. Melo, Aníbal E. Vercesi, and Roger F. Castilho Abstract The mitochondrial electrical membrane potential (Dy) is the main component of the proton motive force (Dp) generated across the inner mitochondrial membrane during electron flow through the respiratory chain. Among the techniques available to assess Dy, methods that rely on the spectrophotofluorometric responses of dyes are widely employed for whole suspensions of isolated mitochondria or permeabilized cells. Safranine is one of the dyes currently used most often for this purpose. Safranine is a lipophilic cationic dye that undergoes optical shifts upon its potential-dependent distribution between the external medium and the intramitochondrial compartment and on its stacking to inner mitochondrial membrane anionic sites. The association between the optical changes of safranine and the membrane potential allows unknown Dy values to be estimated from an equation describing their relationship. Here, we describe the use of safranine as a fluorescent indicator of Dy in isolated mitochondria and digitonin-permeabilized cells. We present suitable conditions to employ safranine as a Dy indicator. Key words: Bioenergetics, Digitonin, Energy metabolism, Membrane potential, Mitochondria, Oxidative phosphorylation, Safranine

1. Introduction The protonmotive force (Dp) across the inner mitochondrial membrane is the energy transduction link between substrate oxidation and mitochondrial ADP phosphorylation (1). FAD and NAD coenzymes are reduced during the degradation of glucose and fatty acids through the glycolytic and b-oxidation pathways, respectively. These pathways also feed acetyl coenzyme A to the Krebs Cycle,

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_7, © Springer Science+Business Media, LLC 2012

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which accomplishes the remaining degradation of the substrates and yields further amounts of reduced FAD and NAD (2). Amino acids are also broken down and fed into the Krebs cycle. The reduced coenzymes, NADH and FADH2, deliver electrons to respiratory chain complexes I and II, respectively. Electrons are then carried from these entry complexes down to complex IV, where molecular oxygen accepts electrons and is reduced to water. Respiratory complexes I, III, and IV couple the electron transport to the pumping of H+ ions from the matrix to the intermembrane space (3). This electron transport through the respiratory chain builds up a resting Dp of approximately −220 mV, which is composed of two thermodynamic potentials: an electrical gradient (Dy) and a pH gradient ( D pH) across the inner mitochondrial membrane (4). The energy stored as Dp drives the re-entry of protons through the ATP synthase complex with concomitant ADP phosphorylation to ATP (5). In energized nonphosphorylating mitochondria, a Dy of nearly −170 mV is by far the largest component of the entire Dp; hence, the contribution of the DpH component is minor. In an experimental setup with isolated mitochondria, the DpH component of Dp can be further decreased by adding permeant anions, if desired, forcing the Dy to increase its own contribution to the Dp (6). Due to the ease of measurement and the representativeness of Dy to the entire Dp, Dy is often the only component measured in cellular and mitochondrial studies that approach questions related to the Dp. The assessment of Dy will depend on the biological sample (i.e., isolated mitochondria, permeabilized cells, and intact cells) and the instrument employed, such as cation selective electrode, spectrophotometer, spectrofluorometer, microscope, and flow cytometer. For the evaluation of Dy in intact cells, the influence of the plasma membrane potential on the measurements should also be taken into account (7). When evaluating whole suspensions of isolated mitochondria or permeabilized cells, the estimation of Dy usually relies on the distribution of membrane permeant cations measured by selective electrodes (8, 9) or the spectrophotofluorometric responses of dyes (10). The dyes currently used most often for this purpose are rhodamine 123 (11) and safranine (12). In our laboratory, over the last two decades, we have employed safranine to estimate Dy in mitochondria isolated from plants (13, 14) and several animal tissues (15–19), and in a variety of permeabilized cells (20–22). Safranine is a lipophilic cationic dye that undergoes optical shifts upon its potential-dependent distribution between the external medium and the intramitochondrial compartment and on its stacking to inner mitochondrial membrane anionic sites (23). The spectral shifts of safranine elicited by the energized states of mitochondria were first described in mitochondrial suspensions by Colonna et al. (24). The almost linear association between safranine absorbance changes and the membrane potential (as titrated

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by increasing extramitochondrial K+ concentrations in the presence of valinomycin) allowed unknown Dy values to be estimated by an equation describing their relationship (12). Later, the fluorescence quenching of safranine upon Dy generation was investigated in vesicles and reconstituted proteoliposomes (25, 26). Monitoring Dy through the fluorescence of safranine allows the use of a lower concentration of the dye, and this may represent the main advantage of fluorescence over absorbance measurements (see Notes 1 and 2). In addition, measurement of the safranine absorbance requires special spectrophotometers that are able to perform dual wavelength measurements simultaneously (12). Here, we describe the features of safranine as a fluorescent indicator of Dy in isolated mitochondria and digitonin-permeabilized cells.

2. Materials 2.1. Isolated Rat Liver Mitochondria

1. Female Wistar Unib Hannover rats were housed at 23°C on a 12-h light–dark cycle with free access to a standard chow diet and water. 2. The following potassium-free isolation buffers were prepared on the day before mitochondrial isolation and stored at 4°C: Isolation buffer I (250 mM sucrose, 1 mM ethylene glycolbis(2-amino-ethylether)-N,N,N ¢,N ¢-tetra-acetic acid (EGTA), and 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer, pH 7.2); isolation buffer II (250 mM sucrose, 0.2 mM EGTA, and 10 mM HEPES buffer, pH 7.2); and isolation buffer III (250 mM sucrose and 10 mM HEPES buffer, pH 7.2).

2.2. PC12 Cell Culture

1. PC12, a cell line derived from a rat adrenal pheochromocytoma, was obtained from ATCC (Rockville, MD, USA). 2. Dulbecco’s modified Eagle’s medium (DMEM) containing 4.5 g/L glucose, with or without phenol red. This medium was supplemented with 10% horse serum, 5% fetal bovine serum, 100 U/mL penicillin, and 100 mg/mL streptomycin. 3. Trypsin solution (0.25%) containing 1 mM ethylenediamine tetra-acetic acid (EDTA), osmolarity 295 ± 5 mOsm/L, and pH 7.0 ± 0.2. 4. Phosphate-buffered saline (PBS), pH 7.2, was prepared weekly and stored at 4°C. 5. A stock solution of digitonin (3.25 mM) was prepared in dimethyl sulfoxide (DMSO) and stored at −20°C. 6. The cell cultures were maintained in CO2 incubator.

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2.3. Determination of the Mitochondrial Membrane Potential in Isolated Organelles or DigitoninPermeabilized Cells

1. Standard reaction medium: 250 mM sucrose, 10 mM HEPES buffer, 200 mM EGTA, 2 mM H2PO4, 1 mM MgCl2, and complex I-linked substrate cocktail (3.4 mM malate, 1.86 mM a-ketoglutarate, 2.1 mM pyruvate, 2.1 mM glutamate), pH 7.2, at 28°C. For the experiments with PC12 permeabilized cells, this medium was supplemented with 0.1% bovine serum albumin. 2. Hyposmotic reaction medium: the same composition was used as for the standard reaction medium described above, except that 50 mM sucrose was used instead of 250 mM sucrose. These reaction mediums were prepared on the day before use and frozen until use. 3. Stock solutions: 2.5 mM safranine in water (see Note 3); 40 mg/mL valinomycin in DMSO; 1 M KCl in water; 50 mM ADP in water (the pH was corrected with NaOH to 7.2); 2 mg/mL oligomycin in ethanol; 1 mM carboxyatractyloside (CAT) in water; 1 mM carbonyl cyanide p-trifluorophenylhydrazone (FCCP) in ethanol; and 5 mg/mL alamethicin in ethanol. All of these reagents were stored at −20°C for up to 6 months. 4. A spectrofluorometer equipped with magnetic stirring and a temperature-controlled water bath was used. The following settings were used: high sensitivity, 2 Hz acquisition rate, excitation wavelength 495 nm, emission wavelength 586 nm, and both slits were set to 5 nm (see Note 4). The temperature was set at 28°C in all experiments. 5. A 3-mL cuvette was equipped with a magnetic stirring bar. All of the experiments were carried out with continuous stirring.

3. Methods The cationic dye safranine is distributed across the inner mitochondrial membrane (negative inside) according to the Dy. The decay in intensity of the safranine fluorescence upon mitochondrial energization arises from the electrophoretic transportation of this dye from the extramitochondrial to the intramitochondrial compartment. The intramitochondrial accumulation of safranine in the millimolar range and its stacking to the inner mitochondrial membrane anionic sites are accompanied by spectral changes and fluorescence self-quenching due to the aggregation of safranine into dimers and polymers (23, 27). Therefore, the dye sample (protein) ratio may greatly affect the fluorescence response (26, 28). The purpose of the experiments shown below was to describe the response of the safranine fluorescence to different experimental conditions, as the safranine and mitochondrial concentrations were

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varied in the suspension. In addition, the response of the safranine fluorescence to different densities of permeabilized cells was studied. In this way, suitable conditions to employ safranine as a Dy indicator were determined. 3.1. Determination of the Mitochondrial Membrane Potential in Isolated Organelles

1. Mitochondrial Isolation: Mitochondria were isolated by differential centrifugation as described by Kaplan and Pedersen (29). All of the procedures after excision of the liver were performed either on ice or at 4°C. Briefly, 2-month-old fed rats were decapitated, and the livers were immediately removed and stored in isolation buffer I. After two washes with isolation buffer I, the liver was finely diced with scissors. Approximately 10 g of tissue was then transferred to the homogenizing potter (#23, Kontes Glass Co. Vineland, NJ), which was filled to 40 mL with isolation buffer I. The tissue was thoroughly homogenized (usually within about ten strokes) while the homogenizing potter was kept on ice. This homogenate was transferred to 40-mL-lidded tubes and centrifuged at 800 × g for 10 min. The supernatant thus obtained was then transferred to a clean tube while avoiding contamination with the loose pellet. This tube was filled completely with isolation buffer I and centrifuged at 7,750 × g for 10 min. After this, the supernatant was discarded and the pellet was suspended in isolation buffer II by gentle mixing with a small paintbrush and transferred to a clean tube. This tube was filled with isolation buffer II and centrifuged again at 7,750 × g for 10 min. The supernatant was discarded and the pellet was washed by rinsing once with 400 mL of isolation buffer III. This pellet was then carefully resuspended in 400 mL of isolation buffer III by gently mixing with a small paintbrush. Finally, this suspension was transferred to a clean glass tube and kept on ice until use. The protein concentration of this suspension was determined using a modified Biuret assay. 2. Membrane Potential Experiments: The fluorescence reading (in the time course mode) was started with standard reaction buffer (2 mL) supplemented with 5 mM safranine in the cuvette (see Note 5). After a short base line reading was recorded, mitochondria were added to the cuvette to a final concentration of 0.5 mg/mL (Fig. 1a). After the initial fast fluorescence decay and the establishment of a steady-state signal, the potassium ionophore valinomycin (40 ng/mL) was added. Thereafter, KCl was consecutively added to clamp the Dy at decreasing values. The first three KCl additions were 0.375 mM each, the next three were 0.5 mM each, and the following four were 0.75 mM each. Between the KCl additions, a steady-state of the fluorescence was waited to re-establish. After the last KCl addition, the protonophore FCCP was added to dissipate the Dy.

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3. The sums of added potassium at each step (0.375, 0.750, 1.125, 1.625, 2.125, 2.625, 3.375, 4.125, 4.875, and 5.625 mM) were used to calculate the Dy according to the Nernst equation: Dy = 60 × log(KIN+/KOUT+), where KIN+ is the intramitochondrial potassium concentration and KOUT+ is the sum of added potassium at each step. KIN+ was assumed to be 120 mM (see Notes 6 and 7). Although a calibration curve can be built with the absolute fluorescence intensity of each step and its respective calculated Dy, comparisons among different conditions (as detailed below)

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Fig. 1. Estimation of the mitochondrial transmembrane electrical potential and the effects of varying the safranine or mitochondrial protein concentration on the safranine fluorescence responses. (a) A representative trace of the changes in safranine fluorescence in response to titration of the electrical potential with KCl additions in the presence of 40 ng/mL valinomycin (V). The effect of 1.25, 2.5, 5, and 10 mM safranine on the fluorescence response was studied at different concentrations of mitochondrial protein (Mito; 0.125, 0.5, and 1.0 mg/mL, presented in panels b, c, and d, respectively). Calibration curves were plotted as “DF/ F ” as a function of the calculated values of Dy, where “F ” is the fluorescence after FCCP addition and “DF ” is “F ” minus the steady-state fluorescence after each KCl addition. The data was fitted to a second-order polynomial equation (solid line), and “X ” represents the estimated resting membrane potential. The data shown in the figure is representative of three independent experiments. a.u. arbitrary units.

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require that the absolute fluorescence be normalized with respect to the amplitude of fluorescence response under each of these conditions. To do so, we calculated DF/F, where F is the fluorescence intensity after the addition of FCCP and DF is F minus any given fluorescence intensity. Following this, DF/F was plotted as a function of the calculated Dy and fitted to a secondorder polynomial equation (Microcal Origin 8.0) (Fig. 1c, filled circle line). 3.2. Effects of Varying the Concentrations of Mitochondrial Protein and Safranine on the Fluorescence Response

The membrane potential calibration experiment was carried out exactly as described above, using varying concentrations of mitochondria (0.125, 0.5, and 1.0 mg/mL) and safranine (1.12, 2.5, 5, and 10 mM) in the suspension. The calibration curves obtained at each mitochondria and safranine concentration combination are plotted in Fig. 1b–d. The results indicate that the sample protein to safranine concentration ratio can greatly affect the safranine fluorescence response. For example, the fluorescence when using too little safranine (1.12 mM) was not responsive to changes in Dy when the concentration of mitochondrial protein is high (1.0 mg/mL), but it may respond to changes in Dy in the high range (−120 to −170 mV) if the concentration of mitochondrial protein is decreased to 0.12 mg/mL. The fluorescence response appears most reliable over the three mitochondrial protein concentrations studied (0.12, 0.5, and 1.0 mg/mL) at a safranine concentration of 5.0 mM, although 2.5 mM safranine may present reliable responses if the mitochondrial protein is £0.5 mg/mL. Using 5.0 mM safranine, the estimated resting Dys are very close to each other, at −161, −163, and −159 mV, for mitochondrial protein concentrations at 0.12, 0.5, and 1.0 mg/mL, respectively.

3.3. Effects of Mitochondrial Volume on Membrane Potential Measurements

The commonly used technique of tetraphenylphosphonium (TPP+) distribution across the inner mitochondrial membrane to follow the Dy relies on the assumption of a given intramitochondrial volume (30). The volume is necessary to calculate the intramitochondrial TPP+ concentration. However, under certain circumstances, mitochondria can undergo extensive changes in volume (31, 32). This issue regarding the mitochondrial volume precludes the use of the TPP+ technique in some experimental setups. It is also known that changes in the mitochondrial volume affect light scattering (33), hence, the fluorescence intensity of the entire suspension might also be affected. To investigate whether changes in mitochondrial volume affect the fluorescence of safranine, the following experiments were performed: 1. The extent of mitochondrial swelling induced by a hyposmotic medium (20% of the standard medium osmolarity) was assessed by light scattering (excitation wavelength 540 nm, emission

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wavelength 540 nm, excitation slit 1.5 nm, and emission slit 3.0 nm; low sensitivity was used). After the mitochondria were added (0.5 mg/mL final concentration) and the light scattering leveling off, 10 mg/mL alamethicin was added to elicit maximal mitochondrial swelling (Fig. 2a). 2. The mitochondria were incubated in standard and hyposmotic reaction mediums supplemented with 5 mM safranine (Fig. 2b). After the mitochondria were added (0.5 mg/mL final concentration) and a new steady-state in fluorescence intensity had been established, the phosphorylation state was induced by adding ADP (300 mM) and later inhibited using oligomycin (2 mg/mL). FCCP (100 nM) was added at the end of the experiment to dissipate the Dy. 3. After the addition of FCCP, it is observed slightly higher safranine fluorescence values under isosmotic conditions than in an hyposmotic medium. To induce similar mitochondrial volumes under these experimental conditions, 10 mg/mL alamethicin was added to the suspensions incubated under both conditions. The data presented in Fig. 2 shows that the extensive mitochondrial swelling that occurs in response to the hyposmotic medium (Fig. 2a) only decreases by less than 10% the total amplitude of the safranine fluorescence response, i.e., the difference between the fluorescence measured before and after FCCP addition (Fig. 2b). After alamethicin was added to induce maximal mitochondrial swelling, the slight difference in safranine fluorescence between hyposmotic and isosmotic conditions was minimized (Fig. 2b).

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1. PC12 cell culture: Cultured PC12 cells were maintained at 37°C and 5% CO2 in DMEM containing phenol red and supplemented with 10% horse serum, 5% fetal bovine serum, 100 U/mL penicillin, and 100 mg/mL streptomycin. Because PC12 cells present a doubling time of 48–72 h, the cells were passaged every 2 days and plated in 150-cm2 tissue culture flasks at an initial density of 0.5 × 105 cells/cm2. 2. PC12 cell suspensions: After 2 days in culture, PC12 cells from one flask were washed once in PBS and then incubated with 4 mL of trypsin-EDTA solution at 37°C for approximately 2 min. Supplemented DMEM (10 mL) was added and the cells were resuspended by pipetting them slowly up and down. The cell suspension was centrifuged at 2,800 × g for 3 min in a 15-mL conical tube at room temperature. The supernatant was discarded and the cell pellet was resuspended in 5 mL of supplemented DMEM without phenol red, but containing 20 mM HEPES (pH 7.2). The cell number was estimated using a Neubauer chamber (see Note 8). Thereafter, the cell suspension was centrifuged at 2,800 × g for 3 min; the resulting pellet was resuspended to a final density of 40 × 106cells/mL and aliquoted into 200 mL portions (8 × 106 cells each). These aliquots were maintained in 0.6 mL microcentrifuge tubes at room temperature for a maximum of 1.5 h. 3. PC12 cell permeabilization: For the experiments, each cell aliquot was centrifuged at 1,500 × g for 3 min at room temperature using a microcentrifuge, the supernatant was discarded and the pellet was washed once in standard reaction medium. The cell pellet was resuspended in 1 mL of standard reaction medium containing 40 mM digitonin, and incubated for 5 min at 28°C (see Note 9). Digitonin was used to selectively permeabilize the plasma membrane (34). Subsequently, the cell suspension was centrifuged at 1,500 × g for 3 min and resuspended in standard reaction medium to a final volume of 50 mL (see Note 10). 4. Mitochondrial membrane potential measurements in permeabilized PC12 cells: The fluorescence reading (in time course mode) was started with standard reaction medium containing 0.1% BSA, 40 mM digitonin, and 5 mM safranine (see Note 5). After a short base line reading was recorded (Fig. 3a), 50 mL of the cell suspension (PC12) was added to the cuvette to obtain 4 × 106 cells/mL in a final volume of 2 mL. After the fast fluorescence decay and the establishment of a steady-state signal, 40 ng/mL valinomycin was added. Thereafter, KCl was added consecutively. The first three KCl additions were 0.375 mM each, the next four were 0.5 mM each, and the final seven were 0.75 mM each. After the last KCl addition, the protonophore FCCP was added to dissipate the Dy. 5. Calibration curve: The sums of the added potassium at each step (0.375, 0.750, 1.125, 1.625, 2.125, 2.625, 3.125, 3.875,

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4.625, 5.375, 6.125, 6.875, 7.625, and 8.375 mM) were used to calculate the Dy according to the Nernst equation, assuming KIN+ to be 120 mM ((32), also see Notes 6 and 7). As described for isolated mitochondria (see Subheading 3.1, step 3), a calibration curve was built by plotting the DF/F as a function of the calculated Dy. Then, the curve was fitted to a polynomial equation (Fig. 3c, filled triangle line).

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6. To evaluate the oxidative phosphorylation capability of permeabilized PC12 cells and the reliability of the methodology in detecting the expected changes in mitochondrial membrane potential, the experiment shown in Fig. 3b was conducted. After the cell suspension was added (to a final density of 4 × 106 cells/mL) and a new steady-state in fluorescence intensity had been established, the phosphorylation state was induced by adding 400 mM ADP and later inhibited using 10 mM CAT. FCCP (1 mM) was added at the end of the experiment to dissipate the Dy. The decrease in Dy elicited by ADP (shown in Fig. 3b) is valuable as a measure of both: (1) proper cell permeabilization, as ADP is not permeable through the cell membrane; and (2) the ability of the cell mitochondria to phosphorylate ADP to ATP and therefore their functionality. 3.5. Effects of Varying the Concentrations of Permeabilized PC12 Cells on the Fluorescence Response

1. The membrane potential calibration experiment was carried out using 0.5 × 106, 1 × 106, 4 × 106, or 12 × 106 PC12 cells/mL. To obtain these cell densities, different volumes of permeabilized cell suspension (6.2, 12.5, 50, and 150 mL) were added to the standard reaction medium containing 0.1% BSA, 40 mM digitonin, and 5 mM safranine, to a final volume of 2 mL. 2. Membrane potential calibration experiments were carried out as described above (see Subheading 3.4, step 5) using consecutive additions of KCl. The calibration curves obtained at each density of permeabilized PC12 cells are plotted in Fig. 3c. The best fitting curve between second- or third-order polynomial equations was considered (Microcal Origin 8.0). The data presented in Fig. 3c shows that the pattern of safranine fluorescence responses to changes in Dy is dependent on the density of cells in the suspension. Apart from the likely influence of mitochondrial concentration, as shown in Fig. 1b–d, the effects of cell density on the turbidity of the whole suspension may also contribute to the observed responses in permeabilized cells. The estimated resting Dy was −157, −159, −158, and −158 mV at 0.5 × 106, 1.0 × 106, 4 × 106, and 12 × 106 cells/mL, respectively. Despite a very similar resting Dy estimated among the conditions, the densities of 1 × 106 and 4 × 106 cells/mL presented better responses to changes in Dy over the entire range evaluated.

4. Notes 1. Our group previously showed that safranine affects mitochondrial responses to Ca2+-induced mitochondrial permeability transition and to oxidative stress ( 15, 16 ) . These effects are concentration-dependent and clearly evident when the concentration of safranine is >5 mM. Figure 4 presents data

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derived from our previous studies showing the stimulatory effects of safranine on Ca2+ plus phosphate-induced mitochondrial permeability transition – as monitored by the swelling rate (15). Conversely, safranine inhibits Fe2+-citrate-induced mitochondrial lipid peroxidation (16). 2. Safranine at high concentrations (³40 mM) was shown to inhibit rat liver mitochondrial respiration and energy transfer. ADP- and FCCP-stimulated respiration rates were progressively inhibited by safranine above 40 mM, whereas state IV respiration was slightly stimulated by safranine at concentrations higher than 70 mM (23). Safranine (³10 mM) may also slow down the heart mitochondrial Ca2+ uptake rate to a small extent (35). 3. Safranine stock solutions should be protected from light. We usually store safranine stock solutions at −20°C for up to 6 months. 4. The use of a microplate reader to measure safranine fluorescence in the suspension could be considered. However, the need for efficient stirring of the suspension may preclude the use of this instrument. 5. Small variations in the fluorescence baseline, as are usually observed when safranine is added to the cuvette separately for each experiment, can be avoided by instead incorporating safranine into the stock reaction medium used for all the experiments. The reaction medium containing safranine should be protected from light. No change in safranine fluorescence was noted when the reaction medium was maintained at 28–37°C for up to 6 h.

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6. The mitochondrial membrane potential is calculated according to the Nernst equation: Dy = 60 × log(KIN+/KOUT+), where KIN+ is the intramitochondrial potassium concentration and KOUT+ is equal to the added potassium. KIN+ is usually assumed to be 120 mM (e.g., Figs. 1 and 3 and ref. 20). The advantage of this method is that an error in the estimate of KIN+ would cause only a small error in the estimated membrane potential. If the KIN+ were in fact 100 mM, Dy would be overestimated by only 5 mV (12). KIN+ can also be measured by accessible techniques if desired (32). 7. During the safranine-Dy calibration experiment, some degree of mitochondrial swelling may occur due to K+ uptake. Nevertheless, this is not expected to result in an important variation in the intramitochondrial K+ concentration because one compensates for the other. Even though it might comprise a source of error in the estimation of Dy, the calibration protocol is rather valuable in assuring that the safranine fluorescence is responding properly to the changes in Dy. 8. If cell aggregation is observed, 0.01% DNAse may be added during the cell trypsinization and washing steps. Cell aggregation can result from the presence of dead cells in the suspensions. 9. The permeabilization protocol can differ among cell types. Different cells may require different concentrations of digitonin. Digitonin, when used below the optimal concentrations, may lead to an undersized initial safranine response due to incomplete cell membrane permeabilization. Cell staining with trypan blue may be useful in evaluating the extent of membrane permeabilization by digitonin. Digitonin should be avoided in excess as it may lead to a loss of Dy over time due to some degree of mitochondrial membrane permeabilization (36). 10. For calibration purposes, the cells must be washed to remove the cytosolic K+ after digitonin permeabilization. In our experience, the higher the cell density, the greater the degree of unwashed cytosolic K+ remaining in the suspensions.

Acknowledgments The authors are currently supported by grants from the following Brazilian agencies: Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP), Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), and Instituto Nacional de Obesidade e Diabetes. T.R.F. and D.R.M. are graduate students supported by FAPESP fellowships.

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References 1. Mitchell P (1961) Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191:144–148 2. Holmes FL (1993) Hans Krebs: vol 2: Architect of intermediary metabolism, 1933–1937. Oxford University Press, Oxford 3. Mitchell P, Moyle J (1967) Respiration-driven proton translocation in rat liver mitochondria. Biochem J 105:1147–1162 4. Mitchell P, Moyle J (1969) Estimation of membrane potential and pH difference across the cristae membrane of rat liver mitochondria. Eur J Biochem 7:471–484 5. Reid RA, Moyle J, Mitchell P (1966) Synthesis of adenosine triphosphate by a protonmotive force in rat liver mitochondria. Nature 212:257–258 6. Nicholls DG (1974) The influence of respiration and ATP hydrolysis on the proton-electrochemical gradient across the inner membrane of rat-liver mitochondria as determined by ion distribution. Eur J Biochem 50:305–315 7. Nicholls DG, Ward MW (2000) Mitochondrial membrane potential and neuronal glutamate excitotoxicity: mortality and millivolts. Trends Neurosci 23:166–174 8. Kamo N, Muratsugu M, Hongoh R, Kobatake Y (1979) Membrane potential of mitochondria measured with an electrode sensitive to tetraphenyl phosphonium and relationship between proton electrochemical potential and phosphorylation potential in steady state. J Membr Biol 49:105–121 9. Serviddio G, Sastre J (2009) Measurement of mitochondrial membrane potential and proton leak. Methods Mol Biol 594:107–121 10. Waggoner AS (1979) Dye indicators of membrane potential. Annu Rev Biophys Bioeng 8:47–68 11. 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 850:436–448 12. Åkerman KE, Wikström MK (1976) Safranine as a probe of the mitochondrial membrane potential. FEBS Lett 68:191–197 13. Costa AD, Nantes IL, Jezek P, Leite A, Arruda P, Vercesi AE (1999) Plant uncoupling mitochondrial protein activity in mitochondria isolated from tomatoes at different stages of ripening. J Bioenerg Biomembr 31:527–533 14. Fortes F, Castilho RF, Castiti R, Carnieri EG, Vercesi AE (2001) Ca2+ induces a cyclosporin-

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A-insensitive permeability transition pore in isolated potato tuber mitochondria mediated by reactive oxygen species. J Bioenerg Biomembr 33:43–51 Valle VGR, Pereira-da-Silva L, Vercesi AE (1986) Undesirable feature of safranine as a probe for mitochondrial membrane potential. Biochem Biophys Res Commun 135:189–195 Castilho RF, Pereira RS, Vercesi AE (1996) Protective effect of safranine on the mitochondrial damage induced by Fe(II)citrate: comparative study with trifluoperazine. Eur J Drug Metab Pharmacokinet 21:17–21 Bassani RA, Fagian MM, Bassani JW, Vercesi AE (1998) Changes in calcium uptake rate by cardiac mitochondria during postnatal development. J Mol Cell Cardiol 30:2013–2023 Maciel EN, Vercesi AE, Castilho RF (2001) Oxidative stress in Ca2+-induced membrane permeability transition in brain mitochondria. J Neurochem 79:1237–1245 Bento LM, Fagian MM, Vercesi AE, Gontijo JA (2007) Effects of NH4Cl-induced systemic metabolic acidosis on kidney mitochondrial coupling and calcium transport in rats. Nephrol Dial Transplant 22:2817–2823 Vercesi AE, Bernardes CF, Hoffmann ME, Gadelha FR, Docampo R (1991) Digitonin permeabilization does not affect mitochondrial function and allows the determination of the mitochondrial membrane potential of Trypanosoma cruzi in situ. J Biol Chem 266:14431–14434 Oliveira KA, Zecchin KG, Alberici LC, Castilho RF, Vercesi AE (2008) Simvastatin inducing PC3 prostate cancer cell necrosis mediated by calcineurin and mitochondrial dysfunction. J Bioenerg Biomembr 40:307–314 Fernandes MP, Inada NM, Chiaratti MR, Araújo FF, Meirelles FV, Correia MT, Coelho LC, Alves MJ, Gadelha FR, Vercesi AE (2010) Mechanism of Trypanosoma cruzi death induced by Cratylia mollis seed lectin. J Bioenerg Biomembr. doi:10.1007/s10863-010-9268-9 Zanotti A, Azzone GF (1980) Safranine as membrane potential probe in rat liver mitochondria. Arch Biochem Biophys 201:255–265 Colonna R, Massari S, Azzone GF (1973) The problem of cation-binding sites in the energized membrane of intact mitochondria. Eur J Biochem 34:577–585 Singh AP, Nicholls P (1984) Energized transport of potassium ions in the absence of valinomycin by cytochrome c oxidase-reconstituted vesicles. Biochim Biophys Acta 777:194–200

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26. Singh AP, Nicholls P (1985) Cyanine and safranine dyes as membrane potential probes in cytochrome c oxidase reconstituded proteoliposomes. J Biochem Biophys Methods 11: 95–108 27. Bunting JR, Phan TV, Kamali E, Dowben RM (1989) Fluorescent cationic probes of mitochondria: metrics and mechanism of interaction. Biophys J 56:979–993 28. Perevoshchikova IV, Sorochkina AI, Zorov DB, Antonenko YN (2009) Safranine O as a fluorescent probe for mitochondrial membrane potential studied on the single particle level and in suspension. Biochemistry (Mosc) 74:663–671 29. Kaplan RS, Pedersen PL (1983) Characterization of phosphate efflux pathways in rat liver mitochondria. Biochem J 212:279–288 30. Kamo N, Muratsugu M, Hongoh R, Kobatake Y (1979) Membrane potential of mitochondria measured with an electrode sensitive to tetraphenyl phosphonium and relationship between proton electrochemical potential and phosphorylation potential in steady state. J Membr Biol 49:105–121

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31. Halestrap AP (1989) The regulation of the matrix volume of mammalian mitochondria in vivo and in vitro and its role in the control of mitochondrial metabolism. Biochim Biophys Acta 973:355–382 32. Kowaltowski AJ, Castilho RF, Vercesi AE (2001) Mitochondrial permeability transition and oxidative stress. FEBS Lett 495:12–15 33. 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 34. Fiskum G, Craig SW, Decker GL, Lehninger AL (1980) The cytoskeleton of digitonintreated rat hepatocytes. Proc Natl Acad Sci USA 77:3430–3434 35. Harris EJ, Baum H (1980) Uptake of safranine by cardiac mitochondria. Competition with calcium ions and dependence on anions. Biochem J 192:551–557 36. Hoppel C, Cooper C (1968) The action of digitonin on rat liver mitochondria. The effects on enzyme content. Biochem J 107:367–375

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Chapter 8 Fluorescence Measurement of Mitochondrial Membrane Potential Changes in Cultured Cells David G. Nicholls Abstract The mitochondrial membrane potential is the dominant component of the proton-motive force that is the potential term in the proton circuit linking electron transport to ATP synthesis and other energy-dependent mitochondrial processes. Cationic fluorescent probes have been used for many years to detect gross qualitative changes in mitochondrial membrane potentials in intact cell culture. In this chapter, I describe how these fluorescence signals may be used to obtain a semiquantitative measure of changes in mitochondrial membrane potential. Key words: Mitochondria, Membrane potential, Neuron, Glutamate, Proton-motive force, Proton electrochemical potential, Tetramethylrhodamine methyl ester, Rhodamine 123, JC-1

1. Introduction The chemiosmotic proton circuit links the mitochondrial electron transport chain to the ATP synthase and other pathways (proton leaks, nicotinamide nucleotide transhydrogenase, metabolite transporters, etc.) that utilize the proton current generated by the proton-translocating complexes I, III, and IV (1). The proton electrochemical potential of the proton gradient across the inner membrane is given (in millivolts) by the proton-motive force (Δp) Dp = Dy - 61DpH at 37°C, where Δψ is the membrane potential (i.e., the difference in electrical potential between the matrix and cytoplasm) in millivolts and ΔpH is the pH difference across the inner membrane. Under most physiological conditions, the contribution of the membrane potential to

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_8, © Springer Science+Business Media, LLC 2012

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the total Δp is dominant (roughly 150 mV) and the ΔpH of typically −0.5 units contributes about 30 mV. Absolute determinations of the components of Δp in intact cells are exceedingly complex (2) and reliant on multiple assumptions, and virtually all studies not only focus on changes rather than absolute measurements but also do not consider the ΔpH component of Δp. Even with these simplifications in mind, the determination of relative changes in mitochondrial membrane potential is far from trivial. An extremely wide range of cells and experiments are performed where mitochondrial membrane potential changes are monitored. For the purposes of this chapter, I shall focus on a single cell type, neurons cultured from rat cerebellum, and detail how semiquantitative measurements may be made of changes in mitochondrial membrane potential (Δym) under a range of conditions using fluorescent membrane-permeant cations (so-called “mitochondrial membrane potential indicators”). An important proviso is that the fluorescence of these indicators is affected by changes in plasma membrane potential and the consequences of this are discussed. Interpretation of the fluorescence traces is not trivial, particularly if there are indications that Δyp is changing during the experiment (following for example activation of plasma membrane ion channels). For that reason, we developed an Excel spreadsheet to deconvolute the traces in terms of dynamic changes in both Δyp and Δym (3). Access to this is described in the text. More recently, the technique has been expanded to include an independent anionic indicator of plasma membrane potential in parallel with the cationic “mitochondrial” indicator (4). The constraints of this chapter do not permit a detailed description of this additional technique, which can be accessed online in the paper and accompanying supplementary material. This semiquantitative technique is currently being refined to improve quantification (Gerencser, Nicholls, and Brand, unpublished). The approach taken in this chapter is not unique; additional programs to interpret fluorescent membrane potential traces exist (5, 6).

2. Materials Unless otherwise stated, all chemicals may be purchased from Sigma-Aldrich and fluorescent probes from Invitrogen-Molecular Probes. Disposables are from VWR or similar suppliers. 2.1. Cell Culture and Incubation Media

(It is assumed that readers are familiar with the conditions required to culture their cells of interest). 1. Culture chambers: Labtech 8-well chambered Cover glasses, sterile (NUNC) VWR product # 43300–774. The wells are previously coated with 33 microg/ml polyethyleneimine and allowed to dry overnight.

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2. Ambient CO2: For microscopes that do not have the ability to maintain a 5% CO2 environment during the experiment. 3.5 mM KCl, 120 mM NaCl, 1.3 mM CaCl2, 1.2 mM MgCl2, 0.4 mM KH2PO4, 5 mM NaHCO3, 1.2 mM Na2SO4, 15 mM D-glucose, 20 mM Na-TES, pH 7.4 at 37°C (see Notes 1 and 2). 2.2. Fluorescent Indicators (See Note 3)

1. Tetramethylrhodamine methyl ester (TMRM perchlorate, Invitrogen, T-668) is dissolved in DMSO at a concentration 100–500 times greater than the final concentration to be used in the assay (which is typically 2–100 nM depending on mode, see below) and is protected from light. Cells in incubation medium are equilibrated with the probe for 45–60 min. In contrast to R123 the experiment is performed without washing away the probe. 2. Rhodamine 123 (R123, Invitrogen R-302) is dissolved in DMSO at typically 2.6 micromolar final concentration. Cells in incubation medium are equilibrated with the probe for 15 min at 22°C, and are washed with fresh incubation medium not containing R123. In contrast to TMRM, the probe is not added to the experimental medium.

2.3. Fluorescence Imaging

1. For confocal imaging, the minimal requirements are for an inverted microscope equipped with a 20× objective with argon and HeNe lasers capable of excitation at 488 nm (or 514 nm) and 543 nm. For single probe imaging, simple long-pass (LP) emission filters may be used. Other possibilities of course exist. 2. A computer controlled stage is an advantage in that, if the microscope’s software allows (as in the Zeiss equipped with the Multitimer option), fields can be defined in up to eight separate wells of the LabTek chamber allowing up to eight longterm experiments to be performed in parallel. 3. Autofocus control is important in long-term experiments. With suitable attenuation, the autofocus configuration can be set to detect the reflected light from the glass–sample interface and then to move a predetermined distance into the sample to optimize the focal plane. 4. For short-term (<30 min) experiments, single field time-course imaging (rather than the MultiTimer option of the Zeiss confocal) is to be preferred to allow greater time resolution. 5. Stringent confocality is not important when single-cell (rather than single mitochondrial) resolution is employed. Indeed with the 20× 0.95 NA air objective employed for these studies, a sufficiently wide pinhole is used to allow a 5–10 nm depth of focus. Wide-field imaging with suitable excitation and emission filters is equally suitable for most studies.

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2.4. Temperature Control

1. Accurate temperature regulation is critical for metabolic studies of membrane potentials. The best solution is to enclose the entire microscope above the focus controls within a temperature controlled enclosure that may be obtained commercially for most microscopes. The author has constructed enclosures from acrylic (plexiglass) sheet that allow access through multiple doors to the microscope, with the focus control and the heat generating lamp enclosure outside. 2. Temperature control is obtained with a Warner TC 344B Dual Automatic Temperature Controller with the temperature sensors located close to the stage. Since the controller has a maximum output of 18 W per channel, the output from one channel is amplified via an assembled Velleman (vellemanusa. com) DC Controlled Dimmer K8003 (now replaced by the equivalent K8064) to power a 150 W vivarium ceramic heater. The second channel of the TC 344B is used simply to control three internal fans to ensure air circulation. This assembly maintains the microscope stage and objectives within 0.5°C of the desired temperature. 3. If a full enclosure is not available, conventional heated stage inserts work, but particularly if oil immersion objectives are employed, it is essential to heat the objective to prevent it from acting as a cold sink. This last step should be done cautiously, as frequent warming and cooling of the objective can introduce internal strains in the optics.

3. Methods In this section, I describe a representative set of applications to monitor membrane potential changes in primary neuronal cultures. In order to interpret the resulting, frequently complex, fluorescence traces, it is essential to have some understanding of the basic principles that govern whole cell fluorescence. These have been covered in publications (3, 4, 7) but are restated here. These simple principles allowed us to formulate Excel spreadsheets that model with a surprising degree of accuracy the predicted responses of cells loaded with different concentrations of R123 or TMRM in response to simultaneous changes in Δym and Δyp (see Note 9). Since many experimental protocols involve addition of agents that will alter Δyp (e.g., ionotropic receptor and channel activators), it is essential to be aware of the influence this will have on the fluorescence signal.

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1. Lipophilic cations and anions seek to achieve a Nernstian equilibrium across both the plasma and mitochondrial membranes. The equilibrium concentration of TMRM+ in the cytoplasm (c) and mitochondrial matrix (m) relative to the external medium (e) is given at 37°C by the following: éëTMRM + ùû = éëTMRM + ùû ·10Dy m /61 m c éëTMRM + ùû = éëTMRM + ùû ·10- Dy p /61 c e (Dy m - Dy p )/61 + + ëéTMRM ûù m = ëéTMRM ûù e ·10 It is, therefore, apparent that the accumulation of TMRM in the matrix is equally sensitive to changes in either potential 2. While the probes are nonselectively permeable across lipid bilayer regions of both the plasma and mitochondrial membranes, the far greater surface to volume ratio of the mitochondrial matrix compared to the cell body means that the probes will redistribute across the former in response to a perturbation much faster than across the plasma membrane. For most practical rates of data acquisition, it can be assumed that redistribution across the inner mitochondrial membrane is instantaneous. 3. At a critical concentration in the matrix, the probes undergo aggregation. The aggregated probe is nonfluorescent and the mitochondrial fluorescence therefore saturates. It is critically important to decide in advance whether this aggregation (or quenching) is to be exploited or avoided.

3.2. Preliminary Considerations

1. Does your cell posses a multidrug transporter?: Before performing any experiments it is essential to determine from the literature or by experiment whether the cells possess an MDR that can disturb and invalidate the equilibration of the probe across the plasma membrane. R123 is a common substrate used to assay MDR activity. If activity exists, it must be inhibited by addition of an MDR inhibitor that will not itself alter the metabolic processes that the experiment is designed to investigate. 2. Quench or nonquench?: Quench mode is a sensitive means to detect rapid changes in Δym that occur during the experiment, for example if ATP synthesis is inhibited by oligomycin, or if a protonophore is added. Quench mode does not detect preexisting differences in Δym between two populations (for example comparing two cell types) and incidentally must therefore never be used for flow cytometry. Changes in Δym and Δyp can frequently be distinguished even with a single indicator based on the direction and rapidity of the fluorescent changes. In nonquench mode at single-cell resolution, redistribution of probe from matrix to cytoplasm does not produce a change in signal, and what is detected is the redistribution of probe across the plasma membrane

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to reestablish the Δyp Nernst equilibrium. In this mode, but not for quench mode, it is, therefore, advantageous to use the more permeant TMRM, and also to include 1 μM tetraphenyl boron throughout the protocol (3). The TPB anion accelerates the equilibration of TMRM across the plasma membrane without affecting the final equilibrium. 3. TMRM or R123?: the two probes are structurally related. R123 is more hydrophilic and equilibrates across the plasma membrane about 20 times more slowly than TMRM. For that reason R123 is loaded by brief exposure to a high concentration of probe, followed by washing (in contrast to TMRM which can achieve equilibrium within 60 min). This means that R123 loading is difficult to control and in practice R123 is always used in quench mode. However because of its slower equilibration across the plasma membrane R123 is less sensitive to Δyp changes than TMRM (see Fig. 2). 4. TMRM alone or in combination with an anionic plasma membrane potential indicator (PMPI)?: Because nonquench mode is dependent on equilibration across the plasma membrane, it follows that the single-cell fluorescence is sensitive to changes in both Δyp and Δym. If the experimenter can be confident that one of these is not changing during the experiment, this will not be a problem; otherwise, it may be necessary to perform a dual label experiment with both TMRM and the anionic probe PMPI (4), the main proprietary component of the Molecular Devices “membrane potential assay kit, explorer format” (R-8042). If the microscope possesses suitable filters, this does not pose a technical problem, although it is important to adjust the concentrations of PMPI and TMRM (nonquench mode) to roughly equalize fluorescent intensities to facilitate subsequent deconvolution. This brief chapter is restricted to the use of single cationic indicators (TMRM and R123); full details of the use of PMPI and TMRM in concert is contained in (4), together with an Excel spreadsheet and a description of the underlying mathematics that may be accessed online at the Journal of Biological Chemistry. 5. Other probe combinations: Monitoring of Δym can readily be combined with other functional indicators as long as sufficient spectral resolution permits. An extremely valuable combination in addition to Δym + Δyp is Δym + cytoplasmic free Ca2+ (Fluo4-AM etc.) since this can be used to establish single cell correlations between stochastic changes in both parameters (3). 3.3. Predictive Modeling of Responses

We have published two Excel spreadsheets with the dual goal of helping with both the design and interpretation of in situ mitochondrial membrane potential experiments. The first, simpler spreadsheet (3) is relevant to the present chapter and may be

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obtained from the author (see Note 4). It facilitates interpretation of the fluorescence changes produced by TMRM or R123 in both quench and nonquench modes in response to changes in either membrane potential. In this model Δyp is not determined directly but is inferred from the curve-fitting traces. The second spreadsheet (4) is more complex because it is designed to interpret parallel changes in PMPI and TMRM (in nonquench mode) fluorescence. Full details of the use, calibration, and interpretation of these dual probe experiments can be accessed in the online journal. Either spreadsheet can be used in two ways. In the predictive mode, expected changes in potential are fed into the program which generates a single-cell fluorescence trace approximating to the subsequent experimental trace. This is valuable for designing experiments, deciding on optimal concentrations of probe and reinterpreting existing results in the literature. In the analytical mode, the experiment is first performed and then the computer simulation is adjusted manually to provide a best fit of the experimental trace, the readout being the membrane potential time-course that produces the best fit. To reduce the degrees of freedom, a number of constants have to be determined or assumed (Subheading 3.4). It is important to emphasize that the techniques described in this chapter do not determine the mitochondrial membrane potential, but rather produce a semiquantitative estimate of changes from an initial defined membrane potential. The value for the starting potential can be taken from the very limited attempts to determine an absolute value. Alternatively, this group has tended to start with an arbitrary 150 mV for Δym, since the absolute value of the potential does not affect the interpretation of the changes in potential seen during an experiment. Perhaps more importantly, a “null-point” technique can be incorporated into the experiment to determine whether the mitochondria are net generators or consumers of ATP (Subheading 3.5, step 2). 3.4. Spreadsheet: Single Labeled TMRM or R123 Cells

The spreadsheet utilizes the three basic principles defined in the introduction to Subheading 3.1. A full description of the mathematical assumptions inherent in the calculations is found in the Appendix of ref. 3. The interface is shown in Fig. 1. Starting values are required for Δyp (from the appropriate electrophysiology literature) and Δym (see above), the fractional volume of the cell occupied by the mitochondrial matrix, the first-order rate constant for the reequilibration of the probe across the plasma membrane of cell being investigated and the quench limit. Each of these may be determined experimentally as described below, or for more qualitative purposes the values predetermined for rat cerebellar granule neurons (4) may be used as starting values.

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Fig. 1. Excel spreadsheet interface for the predictive and analytic modeling of the single-cell fluorescence response of TMRM or R123 to changes in Δym or Δyp. The setting of the parameters is described in the text. The red trace models the total cell fluorescence and the black trace the mitochondrial contribution to the total signal. Values in blue may be modified to simulate a range of experiments and conditions. The spreadsheet may be obtained from the author.

1. Determination of the quench limit (Q, the concentration at which probe aggregation is initiated in the mitochondrial matrix): Once starting values for Δym and Δyp are decided, cells are equilibrated with a range of external TMRM concentrations and an imaging time course is initiated for each. 2 μM FCCP is added, and if a transient “spike” in fluorescence is seen followed by a decay (see for example Fig. 2), then the experiment is in quench mode. The experiment is repeated with decreasing TMRM concentrations until the spike is no longer seen (see ref. 4, Fig. 4). The matrix quench limit Q is then calculated from this external probe concentration (TMRM)e Q = [TMRM]e ´ 10(Dyp + Dym)/60 2. Determination of the plasma membrane rate constant V for TMRM: to allow interpretation of dynamic, rather than steadystate, changes in potential it is necessary to establish the rate constant for the equilibration of TMRM across the plasma membrane. Because of differing surface–volume ratios of cells, it is advisable to obtain this data for each cell type investigated, although semiquantitative results can be obtained using a value of V obtained in other cells allowing for altered cell body diameter. In practice, the same experiment that was used to determine Q above can be used to obtain V. Using the spreadsheet, adjust the value of the plasma membrane rate constant V until the decay rate in the simulated fluorescence matches that of the quenched experiments. Some adjustment will be necessary depending on the experimental conditions. From experience, the rate constant for R123 is 10–20-fold slower than for TMRM (3).

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Fig. 2. Single soma fluorescence of cerebellar granule neurons equilibrated with 50 nM TMRM (a) or loaded for 15 min at 22°C with 2.6 μM R123 (c) were exposed to 100 μM glutamate plus 10 μM glycine. Where indicated, 2 μg/ml oligomycin and 1 μM FCCP were added. (b, c) simulated traces fitted to the following potential changes: Δyp, depolarization from −60 to −20 mV on addition of glutamate; Δym, depolarization from 150 to 145 mV on addition of glutamate, hyperpolarization to 155 mV on addition of oligomycin and collapse of potential with FCCP. Data from ref. 3.

3. Fractional matrix volume: the volume fraction x of the soma occupied by the mitochondrial matrix can be estimated by determining the residual cytoplasmic fluorescence after mitochondrial depolarization by calculating the ratio of the wholecell TMRM+ fluorescence (in nonquench mode) for a cell with

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depolarized mitochondria (e.g., in the presence of myxothiazol to inhibit the respiratory chain and oligomycin to block the ATP synthase) relative to the same cell with polarized mitochondria prior to the addition of inhibitors. If the fraction of the cell occupied by the mitochondrial matrices is x, then the ratio for the whole cell fluorescence for depolarized vs. polarized mitochondria, (ΣTMRM+)depol/(ΣTMRM+)pol will be given by:

(STMRM )depol / (STMRM )pol = 1 / (1 + x·10Dy

m

/60

)

The ratio determined for cerebellar granule neurons (4) was 0.12, substituting this value into the above equation gives a value for the volume fraction x of 2.3% when Δym is 150 mV. It must be emphasized that this assumes that there is no potentialindependent binding of TMRM to components of the cell. Allowing for such binding would indicate an increased matrix volume fraction; however the only difference this would make to the simulation of the Δym time-course would be to decrease further the small contribution of the cytoplasmic TMRM to the whole-cell signal. 3.5. Predictive Modeling of a Neuronal Experiment with the Spreadsheet

The spreadsheet is no longer available as a supplement to ref. 3 but may be obtained freely form the author (see Note 4). The sample experiment will investigate the effect of NMDA glutamate receptor activation on cultured cerebellar granule neurons, and will test whether the mitochondria in the exposed cell is still generating ATP. The program can use both quench and nonquench modes and simulate TMRM and R123. The significance of this approach is that it allows changes in Δym and Δyp to be distinguished on the basis of both the direction and the kinetics of the single cell fluorescence response but only in quench mode. 1. In the sample experiment, granule neurons loaded with TMRM in quench mode are exposed to glutamate/glycine to activate their NMDA receptors. An initial slight mitochondrial depolarization results in a transient increase in fluorescence. The parallel plasma membrane depolarization produces a decrease in cell fluorescence with a much slower time-course due the rate constraints of equilibration across the plasma membrane. 2. Addition of oligomycin performs an important role: this “oligomycin null point test” determines whether the mitochondria in the cell are synthesizing ATP at the moment the inhibitor is added, or whether, due to damage or partial uncoupling, they are hydrolyzing cytoplasmic ATP by reversal of the ATP synthase. In the former case, oligomycin will hyperpolarize the mitochondria (decreased cell fluorescence in quench mode), if they are damaged they will further depolarize on addition of oligomycin (increased fluorescence in quench mode), see ref. 8.

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3. Finally FCCP is added to complete depolarization and confirm that the experiment was still in quench mode prior to its addition. (If there is no FCCP spike it is best to repeat the simulation (and hence the actual experiment) at a higher TMRM concentration). 4. Load the spreadsheet into Excel 2003 or 2007. Save a “master copy”. 5. The only editable cells are in blue. The default values are for rat cerebellar granule neurons as follows: mitochondrial volume fraction 0.023 (4), external probe concentration 100 nM (for quench mode, 5 nM may be used for nonquench mode), plasma membrane rate constants 0.02 (TMRM) or 0.001 (R123), quench limit 70 μM. The interval (to set the time range of the “experiment”) is set to 20 s (giving a total time course of 1 h). The program allows for changes in Δym and Δyp at four time points during the simulation. The default values for these neurons are set to −83 mV and −150 mV respectively. Lines 31–219 are the iterative calculations and are not editable. The graph displays the total cell fluorescent signal and the mitochondrial fluorescence. 6. To model the protocol for determining the quench limit, set the final Δym to zero to simulate protonophore addition and adjust the external probe concentration until the “FCCP” spike just disappears. 7. To model the effects of partial mitochondrial depolarization in quench and nonquench modes, set the final Δym to −120 mV and compare a TMRM concentration of 100 and 5 nM. This shows up a very important limitation of quench mode: namely that small changes in Δym produce only a transient single cell signal – the FCCP “spike” rapidly decays. thus quench mode cannot be used to detect a preexisting changes in Δym. For example it is inappropriate for flow cytometry where Δym in two conditions or cell types are to be compared. This is frequently not appreciated. 8. Repeat #7 with both 5 and 100 nM R123 simulated (set Q to 0.001). 9. To demonstrate the ambiguity of nonquench mode to changes in the two potentials, compare the response to Δym depolarization to −120 mV with Δyp depolarization to −40 mV at 5 and 100 nM TMRM. 10. In the sample experimental protocols below for TMRM and R123, it would be predicted that NMDA receptor activation would depolarize the plasma membrane (say to −30 mV). The mitochondria would partially depolarize in response to the increased Ca2+ uptake and ATP turnover (say to −140 mV). The third time point simulates the mitochondrial hyperpolarization in response to oligomycin (Δyp −30 mV, Δym −160 mV) and in the final time point FCCP is used to collapse Δym.

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11. Keeping this membrane potential protocol compare quench and nonquench concentrations of TMRM and R123. Figure 2 shows a actual experiment, which is detailed below and the accompanying simulation (3). 3.6. Sample Experimental Protocol: Calibration and NMDA Receptor Activation of Cerebellar Granule Neurons with TMRM in Quench Mode

This is the actual experiment that was used in the above predictive simulation. 1. Culture neurons at about 100,000–350,000 cells per well into the appropriate number of wells of the 8-well Labtech chambered coverslip. 2. Take LabTech chamber from incubator. Remove culture medium and immediately replace with 400 μl of incubation medium without MgCl2 (for NMDA receptor studies) but containing 50 nM TMRM (see Note 5). Incubate the cells in an air incubator (or the temperature-controlled enclosure of the microscope) for 60 min to allow equilibration of the probe. 3. Insert the Labtech chamber onto the microscope stage (see Note 6). With the binoculars and visible transmission locate a suitable field and focus. 4. Set up a confocal configuration with 543 nm excitation (although 514 or even 488 nm will also work) and a 560 longpass (or similar) emission filter. 512 × 512 resolution is adequate. Ensure that laser power is low (1–2%) and use an amplifier gain that is high but does not produce noise in the image. 5. To make additions during an experiment, pipette the required volume of reagent into a 0.5-ml Eppendorf tube. Equip a 100-μl micropipette with a long gel loading pipette tip (Microflex 0.1–200 μl VWR # 53503–189) whose last 1 cm is bent downward at 45°. Carefully withdraw 50 μl of incubation medium from the well that is being imaged, mix thoroughly with the reagent and smoothly readd to the well, ensuring full mixing by three to four slow up and down strokes of the micropipette. With practice, it is easy to do this without disturbing the cells or the focus. 6. To investigate Δym changes in response to NMDA receptor activation, the following additions may be made during a 60-min experiment: (a) 100 μM glutamate plus 10 μM glycine at 5 min (see Note 7). (b) 2 μg/ml oligomycin at 40 min (see Note 8). (c) 1 μM FCCP at 50 min (see Note 9). 7. Save the time-course and define regions of interest (ROI) around individual cells. Export the resulting time-courses to Excel. 8. Interpretation: While the responses of individual cells vary stochastically, an actual trace from a single cell that survived

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the exposure to glutamate/glycine without undergoing delayed Ca2+ deregulation is shown in Fig. 2a. In Fig. 2b, the simulated trace is shown that was generated using the spreadsheet on the assumptions that glutamate caused an immediate Δyp depolarization from −60 to −20 mV. Simultaneously the mitochondria depolarized from 150 to 145 mV (due to enhanced ATP demand and Ca2+ uptake), (3). The small decrease in signal with oligomycin is consistent with a 10 mV mitochondrial hyperpolarization (showing that they were generating ATP). Finally, the FCCP spike confirms that the cell remains in quench mode. 9. Alternative: selected imaging of mitochondrial-poor (nuclear) regions – if resolution permits the regions of interest can be limited to the nuclear regions of individual cells. TMRM and R123 appear to equilibrate freely between cytoplasm and nucleus. In quench mode, this improves the sensitivity of the technique by removing the (constant) mitochondrial signal. In the simulation, this cytoplasmic signal is given by the difference between the whole cell and matrix traces. 3.7. Sample Experimental Protocol: NMDA Receptor Activation of Cerebellar Granule Neurons with R123 in Quench Mode

Rhodamine 123 is structurally related to TMRM but is more hydrophilic and therefore equilibrates more slowly across the plasma membrane (although redistribution across the mitochondrial membrane is still very rapid). R123 fluorescence is, therefore, more slowly affected by redistribution across the plasma membrane, and this can be useful in simplifying the interpretation. R123 is always used in quench mode and is loaded in a different manner (see below). The R123 experiment is identical apart from the loading and fluorescence wavelengths. 1. R123 loading: Cells are equilibrated with 2.6 μM R123 for 15 min at 22°C after which they are washed in incubation medium in the absence of R123. Trial and error is advised to achieve the optimal loading. Be very aware of the risks of phototoxicity; use minimal laser power and avoid exposing the cells to unnecessary excitation. Remember that no R123 is added after washing or during the experiment. 2. Set up a confocal configuration with 488 nm or 514 nm excitation and a 530 nm long-pass (or similar) emission filter. 512 × 512 resolution is adequate. Ensure that laser power is low (1–2%) and use an amplifier gain that is high but does not produce noise in the image. 3. The identical addition protocol to the TMRM experiment is performed. 4. Interpretation – the response of a representative cell that survived glutamate exposure is shown in Fig. 2c. Note that the slopes of the signal following glutamate and FCCP are much shallower than for TMRM, due to the more hydrophilic nature of R123

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and its consequent slower redistribution across the plasma membrane. The simulated trace (Fig. 2d) was obtained using the spreadsheet with all parameters identical to the TMRM experiment, except that the rate constant V was lowered from 0.02 to 0.001 s−1 (3). 5. Employ spreadsheet 1 as detailed in Subheading 3.4 to generate a curve-fit to the experiment.

4. Notes 1. For experiments investigating NMDA glutamate receptors, MgCl2 is omitted to prevent the voltage-block of the receptor under polarized conditions. 2. To work in an ambient CO2 atmosphere, HCO−3 concentration is lowered to 5 mM and a high buffering capacity is introduced (20 mM Na-Tes, pH 7.4) to minimize alkalinization of the medium as CO2 is slowly lost to the atmosphere. 3. TMRM (or the closely related ethyl ester TMRE) and R123 are the accepted “least bad” indicators, in that they give generally reliable results with little interference with normal mitochondrial function. The seductive JC-1, where the aggregate instead of being nonfluorescent fluoresces red, is inappropriate first because it is not valid to ratio the red fluorescence of the aggregate in the matrix and the green monomer fluorescence in the cytoplasm as is frequently done. Second, the aggregates can fail to redissolve when Δym is decreased. Finally, the very slow permeation of the rather hydrophilic probe across the plasma membrane means that it is possible to be misled that for example mitochondria in thin processes posses a higher Δym than mitochondrial in the cell bodies, when this is simply because insufficient time has been allowed for equilibration into the large cell body. Cyanine dyes such as DiOC6 (3) need to be avoided, since many are potent inhibitors of mitochondrial electron transport. 4. The spreadsheet may be obtained free of charge from the author by e-mailing at [email protected] 5. The concentration of TMRM required to remain in quench mode until the final FCCP depolarization may need to be determined in pilot experiments by experimenting with different concentrations and confirming that addition of FCCP gives a transient increase in fluorescence (dequenching). 6. To minimize evaporation, the LabTech chambers have a loose fitting lid. Taking on and off this lid can disturb the focus

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slightly. To allow four experiments to be performed sequentially without evaporation, drill a 8 mm hole in spare lids such that additions can be made to a single well while the remainder are covered. 7. Glycine is a coactivator of the NMDA receptor. To obtain maximal activation of the receptor, glycine is added and Mg2+ is omitted from the medium. 8. Oligomycin inhibits the mitochondrial ATP synthase. If the mitochondria in a cell are still generating ATP at this stage, oligomycin will cause a hyperpolarization in Δym as the proton reentry will be slowed. Conversely, if the mitochondria are proton leaky or with a compromized electron transport chain, the ATP synthase can reverse and hydrolyze cytoplasmic glycolytic ATP. In this case, oligomycin will depolarize the mitochondria. 9. It is useful to conclude the experiment by adding FCCP to collapse Δym and to confirm that the experiment was performed in quench mode. References 1. Nicholls DG, Ferguson SJ (2002) Bioenergetics, vol 3. Academic, London 2. Hoek JB, Nicholls DG, Williamson JR (1980) Determination of the mitochondrial protonmotive force in isolated hepatocytes. J Biol Chem 255:1458–1464 3. Ward MW, Rego AC, Frenguelli BG, Nicholls DG (2000) Mitochondrial membrane potential and glutamate excitotoxicity in cultured cerebellar granule cells. J Neurosci 20:7208–7219 4. 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 5. Ward MW, Huber HJ, Weisova P, Duessmann H, Nicholls DG, Prehn JHM (2007) Mitochondrial

and plasma membrane potential of cultured cerebellar neurons during glutamate induced necrosis, apoptosis and tolerance. J Neurosci 27:8238–8249 6. Lemasters JJ, Ramshesh VK (2007) Imaging of mitochondrial polarization and depolarization with cationic fluorophores. Methods Cell Biol 80:283–295 7. Nicholls DG, Ward MW (2000) Mitochondrial membrane potential and cell death: mortality and millivolts. Trends Neurosci 23:166–174 8. Rego AC, Vesce S, Nicholls DG (2001) The mechanism of mitochondrial membrane potential retention following release of cytochrome c in apoptotic GT1-7 neural cells. Cell Death Differ 8:995–1003

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Chapter 9 Phenomenological Kinetic and Control Analysis of Oxidative Phosphorylation in Isolated Mitochondria Vilmante Borutaite and Rasa Baniene Abstract Metabolic control analysis provides a quantitative framework for analyzing regulatory properties of enzymes in various metabolic pathways. It has been used for estimation of control parameters of the enzymatic pathways at isolated enzyme, cellular, or whole organism levels. This chapter describes how control and elasticities analysis can be experimentally applied to measure control properties of the components of the oxidative phosphorylation system and how a variant of such analysis – phenomenological kinetic analysis – can be used to investigate the effects of various factors (physiological or pathological) on the system of oxidative phosphorylation in isolated mitochondria. Key words: Mitochondria, Oxidative phosphorylation, Respiration, Membrane potential, Metabolic control analysis

1. Introduction Metabolic control analysis (MCA) is a mathematical tool for studying complex biological systems such as metabolic, signaling, or genetic pathways. MCA quantifies how variables, such as fluxes through the system and metabolite/intermediate concentrations, depend on the system parameters. In particular, it describes how the system-dependent properties, so-called control coefficients, depend on local properties, called elasticities. In other words, MCA allows quantitative determination of the contribution of each component to the overall behavior of the system under consideration. Another important aspect is that knowing the control structure of the system (control coefficients exerted by particular components of the system) allows identification of the components of the system that are particularly important in responding to the effects of external factors on the system. Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_9, © Springer Science+Business Media, LLC 2012

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Theoretical background for MCA comes from early works by Higgins (1) and Kascer (2). More recently, MCA has been reviewed by Fell (3) and Brown (4), providing useful experimental applications of the analysis for studying mitochondrial and cellular bioenergetics. There are two main approaches for determining control structure of the system: 1. The “bottom-up” approach, which measures control coefficients of the individual components (enzymes, transporters, etc.) of the system. 2. The “top-down” (also called elasticity or modular kinetic analysis) approach, which measures control coefficients of the larger blocks of the system such as groups of enzymes connected by a common intermediate (metabolite, proton-motive force in the case of oxidative phosphorylation, etc.). These approaches are complementary to each other and can be used separately or both together depending on the tasks of the study. The first approach is useful when it is necessary to know which particular enzyme controls the flux through the system and how much of the control it exerts. In this approach, control coefficients of the individual enzymes within the system are determined by changing their activities with specific inhibitors (or changing activities/levels of enzymes genetically) and measuring changes in the flux through the system. The second approach is used when a researcher wants to know how control is distributed between different parts (blocks of the enzymes) of the system rather than determining input of individual components. In contrast to “bottom-up” approach, here enzymes are conceptually grouped into a small number of blocks within the system instead of analyzing components individually. This simplifies the complex system, and it becomes much easier to analyze its behavior in general, and particularly in complex physiological situations. In this approach, control coefficients are calculated not from the effects of specific inhibitors on the individual enzymes, but the relative elasticities (sensitivities) of the blocks of the system to their intermediate are measured and from them the control coefficients of the blocks are calculated using theorems of MCA. A simplified version of “top-down” analysis (without quantifying control parameters of the system) is used in so-called phenomenological kinetic analysis. In this approach, kinetic responses of the modules of the system to the changing levels of the connecting intermediate are measured in the absence and presence of some external effector of the system. Changes in this kinetics indicate which of the modules are directly impacted by the effector rather than being indirectly affected by changes in the intermediate levels.

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“Top-down” control (modular kinetic) analysis has been previously used for investigations of oxidative phosphorylation system affected by hormones (5), ischemic insult (6), fatty acids and their derivatives (7), heavy metals (8), etc. Phenomenological kinetic analysis has been applied to investigate which parts of the oxidative phosphorylation system are directly affected by elevated concentrations of calcium (9), or how heart preconditioning with nitric oxide affects this system (10). In this chapter, we introduce basic definitions (theorems) of MCA and provide protocols for “top-down” (modular kinetic analysis) of the oxidative phosphorylation system and for measurement of control coefficient of ATPase in isolated mitochondria (“bottom-up” approach). 1.1. Basic Definitions and Theorems in MCA

As any theory, MCA in its basic form relies on certain assumptions and has a number of limitations: 1. The system which is studied consists of the components connected to each other via common intermediates/metabolites forming an independent unit as a whole. 2. The intermediates between the enzymes within the system are freely diffusible. 3. Only infinitesimal changes are considered by the theory; therefore, it is not possible to make the accurate predictions from the measured coefficients about the effects of the large changes in the system. 4. The behavior of the system can be studied in a steady-state only. 5. The coefficients determined in one condition refer only to that particular condition and will be different when condition change. Flux control coefficients (C) quantify the importance of each step in the pathway (single reaction or a block of reactions) in controlling the flux through that pathway in a steady state. It represents the fractional change in the system property (flux, J ) that would be caused by an infinitesimal fractional change in the activity of the particular enzyme (or a block of enzymes) of the system: CiJ =

¶J j ¶E i

´

Ei Jj

where Jj – steady-state flux of the pathway j; Ei – concentration of enzyme i. Control coefficients are the properties of whole system rather than individual enzymes. Elasticity coefficients, unlike control coefficients, are properties of the individual components/enzymes of the system. They measure how isolated enzymes are sensitive to changes in parameters

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(concentrations of substrates or products of reactions, external effectors, etc.). Elasticity coefficient represents the fractional change in the rate of isolated enzyme (block of enzymes) caused by an infinitesimal fractional change in the effector concentration and is expressed: e ki =

¶vi M ´ k ¶M k vi

where vi – the steady-state rate of enzyme i, which is kinetically isolated from the pathway by holding substrate, product, and concentrations of other external effectors constant; Mk – concentration of effector k. Each enzyme may have as many elasticity coefficients as the number of various parameters that affect the enzyme in the system. The flux control summation theorem states that for a given flux through the system the sum of its control coefficients of all steps is equal to 1 (unity): n

åC

J i

=1

i =1

where n – total number of enzymes in the system. The summation theorem implies that the control of flux can be shared by many enzymes of the system. It also implies that increase in some of the control coefficients in the system requires decreases in the others maintaining the sum of all control coefficients constant (1). The connectivity theorem relates flux control coefficients and elasticities of the steps with common intermediates. It states that for every enzyme in the system that responds to changes in concentration of intermediate X, the flux control coefficients and the elasticity coefficients of the enzymes toward X are related in the following way: n

åC

J i

eix =0

i =1

Using connectivity and summation theorems, it is possible to calculate all control coefficients given the enzyme elasticities. For more detailed reading, see (4, 11). 1.2. General Definition of the System for “Top-Down” Modular Kinetic (Elasticity) Analysis

For the analysis, the system of oxidative phosphorylation is conceptually divided into three subsystems, connected by the common intermediate – the proton-motive force, Δp, so that one subsystem generates and two subsystems consume Δp (Fig. 1). Δp is generated by the respiratory subsystem (R), which consists of the transporters of the respiratory substrates, the enzymes of Krebs cycle, substrate dehydrogenases and complexes of the respiratory chain.

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Substrate

Respiratory subsystem Substrate carriers, dehydrogenases, respiratory chain complexes

ΔΨ

139

Proton leak subsystem Membrane permeability to protons, cation cycling Phosphorylating subsystem ATP/ADP translocator, Pi carrier, ATP synthase

ADP ATP

Creatine

Creatine phosphate

Fig. 1. Scheme of oxidative phosphorylation system in heart mitochondria.

Δp is consumed by the phosphorylating subsystem (P), which consists of ATP synthase, the adenine nucleotide translocator, the phosphate carrier, and any of the reactions that convert ADP to ATP (outside or inside mitochondria). Another subsystem consuming Δp is the membrane permeability subsystem (L) consisting of the passive permeability of the mitochondrial inner membrane to protons and any of the reactions cycling cations and dissipating Δp (we call it as a proton leak subsystem). Δp consists of two components: mitochondrial membrane potential (Δy) and proton concentration gradient (ΔpH), D p = D y - zDpH. In general, to estimate Δp both Δψ and ΔpH should be measured. However, in many experimental settings for investigation of oxidative phosphorylation system in isolated mitochondria using buffers with high phosphate and KCl concentrations, ΔpH is low (see 12, 13). Therefore, in such experimental approaches, ΔpH may be neglected and Δp ~ Dy (see Note 1). The kinetic responses of these subsystems to the changes of the proton-motive force (membrane potential) can be determined experimentally by measuring fluxes through the subsystems (modules): 1. The dependence of the flux through the respiratory subsystem (JR) on Δy can be determined by titrating mitochondrial respiration in state 3 with an inhibitor of the phosphorylating subsystem. Carboxyatractyloside or oligomycin (which inhibit phosphorylation and thus raise Δy) can be used.

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2. The dependence of the flux through the proton leak (JL) subsystem on Δy can be determined under conditions of complete inhibition of phosphorylation in the presence of excess oligomycin. In such case, mitochondrial respiration rate is proportional to the rate of proton leak through the mitochondrial inner membrane, since in the absence of phosphorylation all the protons pumped out by the respiratory chain must return to the matrix via the proton leak. Respiration can be titrated with inhibitors of the respiratory chain such as malonate for succinate oxidation and rotenone or myxothiazole for NAD-dependent respiratory substrates. 3. The dependence of the flux through the phosphorylating subsystem (JP) on Δψ can be determined in a similar way as the flux through the proton leak subsystem, but under conditions of active, phosphorylating mitochondrial respiration (i.e., in the absence of oligomycin and in the presence of ADP). Δy is then titrated with an inhibitor of the respiratory chain (malonate for succinate oxidation, myxothiazole or rotenone for NADdependent substrates) and JP at any given Δy is calculated as the difference between the respiration rate in the absence of oligomycin and its presence because in the steady state in the presence of phosphorylation, the rate of proton efflux is equal to the rate of proton influx due to ATP synthesis: JP = JR - JL. Various steady states of mitochondrial respiration can be adjusted by changing concentrations of creatine and creatine phosphate in the medium when creatine phosphokinase ADPregenerating system is used. Other ADP-regenerating systems (such as glucose/hexokinase, etc.) can be also applied (but see Note 2).

2. Materials 1. Basic incubation medium for heart mitochondria: 110 mM KCl, 50 mM creatine, 2.24 mM MgCl2 (corresponding to 1 mM free Mg2+), 10 mM Tris–HCl, 5 mM KH2PO4, 1 mM EGTA, 10 mM DTT, pH 7.2 (at 37°C). This buffer is used to achieve maximal (corresponding to state 3) rate of mitochondrial respiration. 2. Different intermediate steady-state rates of mitochondrial respiration can be adjusted by varying creatine and creatine phosphate in the medium (while keeping total concentration of them 50 mM), and changing concentration of KCl accordingly for the maintenance of constant ionic strength. To obtain approximately 50% of State 3 respiration rate, a buffer containing 80 mM KCl, 20 mM creatine, 30 mM creatine phosphate, 2.53 mM MgCl2

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(1 mM free Mg2+), 10 mM Tris–HCl, 5 mM KH2PO4, 1 mM EGTA, 10 mM DTT, pH 7.2 (at 37°C) can be used. 3. ADP-regenerating system: 400 IU/mL creatine phosphokinase; prepare freshly. 100 mM ATP dissolved in water and adjusted to pH 7.0 with KOH; aliquots can be stored at −20°C for ~1 week; avoid freezing–thawing. 4. Substrate solutions: 100 mM pyruvate plus 100 mM malate dissolved in water and adjusted pH 7.0 with KOH (or Trizma base); store at −20°C for 1–2 weeks. 1 M succinate dissolved in water and adjusted to pH 7.0 with KOH; can be stored at −20°C for several weeks (see Note 3). 5. Tetraphenylphosphonium (TPP+) solution (10 mM of TPPCl in water). A stock solution can be stored at −20°C for several weeks. Working solution of TPP+ (0.05 mM), dilute 5 μL of stock solution with 995 μL of water. 6. Carboxyatractyloside (0.5 mM in ethanol). Aliquot and store at −20°C. 7. Oligomycin (2 mg/mL in ethanol). Store at −20°C. 8. Rotenone (1 mM in ethanol). Store at −20°C. 9. Valinomycin (0.5 mM in ethanol). Store at −20°C.

3. Methods Here, we provide protocols for determining kinetic dependencies of the modules (the respiratory, proton leak, and phosphorylating subsystems) of the oxidative phosphorylation system on the membrane potential in isolated rat heart mitochondria (Fig. 2). To investigate the effect of an external effector (or some physiological/pathological change) on the system, the overall kinetic responses of the modules of the system to Δψ have to be re-measured in the absence and presence of the effector (or with affected and nonaffected mitochondria). This enables to identify the primary sites of action of the effector and to distinguish them from all secondary, indirect changes. For example, such method has been previously used to determine which parts of the oxidative phosphorylation system are affected by ischemia, calcium overload and heavy metals (7–10, 12, 13). In order to determine how control is distributed within the oxidative phosphorylation system the kinetic response curves and theorems of MCA can be further used to estimate elasticity and control coefficients of the modules.

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Respiratory rate, nmolO/min mg protein

300 250 200

A C

150 D 100 50 0 90

B 100

110

120

130

140

150

160

Membrane potential, mV

Fig. 2. Kinetics of the respiratory, proton leak, and phosphorylating subsystems. Open symbols – kinetic dependences of fluxes through the respiratory subsystem (open circle, A trace), the proton leak subsystem (open square, B trace), and phosphorylating subsystem in state 3 (open triangle, C trace). Dark symbols – kinetic dependence of flux through the phosphorylating subsystem in the intermediate steady-state (filled triangle, D trace).

In this method, respiration and membrane potential of isolated mitochondria are measured simultaneously in a closed, stirred and thermostatically controlled vessel fitted with both a Clark-type oxygen electrode and a tetraphenylphosphonium (TPP+)-selective electrode (see Note 4). Rat heart mitochondria can be isolated as described in ref. 9. The experiments are performed at 37°C. 3.1. Simultaneous Measurement of Oxygen Uptake and Membrane Potential

1. Before the experiment wash the oxygen electrode vessel several times with 70% ethanol and de-ionized water. 2. Add 1.5 mL of the incubation medium to the vessel equipped with an oxygen electrode. 3. Immerse a TPP+-sensitive electrode. The electrode should be filled in with 3 mM KCl before the experiment. 4. Add 4 μL of TPP+ chloride (0.05 mM) and wait for 1–2 min until a stable signal is reached (see Note 5). Then put 2 μL TPP+ and after stabilization of the chart recorder trace add the third and fourth 2 μL aliquots of TPP+ as shown in Fig. 3. Do not exceed 1 μM final concentration of TPP+ when NADdependent substrate oxidation is studied (see Note 6). 5. Add 10 μL of 100 mM pyruvate plus 100 mM malate. 6. Add suspension of heart mitochodria (final concentration of mitochondrial protein 0.5 mg/mL) to the vessel. Mitochondria start to respire and accumulate TPP+ (Fig. 3). Monitor mitochondrial respiration and membrane potential for 2–3 min until steady state is reached.

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Valinomycin 1µL

ATP 10 µL TPP+ 2 µL TPP+ 2 µL

Mitochondria 0.5mg/ml

TPP+ 2 µL

Incubation medium 1.5 mL pyruvate 10 µL malate 10 µL TPP+ 4 µL

100% O2

0% O2

chart recorder scale, mm

Fig. 3. Typical traces of simultaneous measurement of respiration and membrane potential of isolated heart mitochondria.

7. Then add 10 μL of 100 mM ATP to achieve the maximal rate of respiration. 8. Finally add 0.5–1 μL of 0.5 mM valinomycin to dissipate membrane potential and to estimate a non-specific TPP+ binding to mitochondria. 3.2. Calculation of Mitochondrial Respiratory Rates and Membrane Potential from the Recordings of the Oxygen and TPP+ Electrodes

1. Convert recordings of chart recorder (or computer) into values of oxygen concentration and calculate the rate of mitochondrial respiration. The concentration of oxygen in air-saturated medium at 37°C has been estimated to be 422 nmolO/mL. Express the rate of oxygen consumption in nmolO/min per mg of mitochondrial protein. 2. Measure the deflection of TPP+ electrode trace after each TPP+ addition in cm from the baseline. 3. Create the calibration graph: plot the deflections of TPP+ electrode trace against the log of the TPP+ concentration. 4. To measure the external TPP+ concentration at any given electrode signal, measure the deflection from the baseline in cm at the desired steady state. Read off the external TPP+ concentration from the calibration graph. 5. Calculate the mitochondrial membrane potential (Δy) at 37°C by using the equation: Dy = 61.5 ´ log([TPP + ]added - [TPP + ]external ) ´

FTPP+ 0.001 ´ mg protein ´ [TPP + ]external

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where FTPP+–TPP+ binding correction factor; use value of 0.162 for heart mitochondria (see Note 7); and [TPP+]added – total added TPP+ concentration in the medium; [TPP+]external – TPP+ concentration in the medium (not consumed by mitochondria). 6. Average the values of at least three repeats on the same preparation of mitochondria. 3.3. Measurement of Kinetic Dependence of the Proton Leak (JL) Subsystem on Dy (Fig. 4)

1. Perform 1–6 steps described in Subheading 3.1. 2. Add 1–2 μL of oligomycin (1 μg/mg of mitochondrial protein) to the incubation medium to completely inhibit phosphorylating subsystem and to achieve the state 4 mitochondrial respiration rate. 3. Add 10 μL of 100 mM ATP. If phosphorylation is completely inhibited by oligomycin, mitochondrial membrane potential and respiration rate do not change after addition of ATP. 4. When the equilibrium is reached, titrate gradually respiration rate with 1–2 μL of 1 mM rotenone, a respiratory chain inhibitor, until oxygen consumption is completely inhibited (see Notes 8 and 9). 5. Plot respiration rate versus Δy as shown in Fig. 4. This represents kinetics of proton leak. 6. Repeat these steps with an effector added or with mitochondria affected by some pathological (physiological) agent to investigate whether it acts on the proton leak subsystem.

3.4. Measurement of Kinetic Dependence of the Respiratory (JC ) Subsystem on Dy (Fig. 5)

1. Perform 1–7 steps described in Subheading 3.1. 2. When the equilibrium is reached, titrate gradually respiration rate with an inhibitor of Δy-consuming processes by adding 1 μL aliquots of carboxyatractyloside (or oligomycin). After each addition wait until traces of oxygen consumption and TPP+ uptake are linear. Repeat this step until oxygen consumption is completely inhibited (see Note 10). 3. Plot respiration rate versus Δy as shown in Fig. 5. This represents kinetics of the respiratory subsystem. 4. Repeat these steps with an effector added or with mitochondria affected by some pathological (physiological) agent.

3.5. Measurement of Kinetic Dependence of the Phosphorylating (JP ) Subsystem on Dy (Fig. 6)

1. Perform 1–7 steps described in Subheading 3.1. 2. When the equilibrium is reached, titrate gradually respiration rate with an inhibitor of Δy-producing processes by adding 0.5–1 μL aliquots of 0.01 mM rotenone. Wait until oxygen consumption and TPP+ uptake are linear. Repeat this step until oxygen consumption is completely inhibited.

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Respiratory rate, nmolO/min mg protein

100

80

60

40

20 × ×× × 0 100

120

140

160

180

200

Membrane potential, mV Fig. 4. Kinetics of the proton leak subsystem. Rate of respiration and membrane potential were titrated with rotenone in the presence of excess of oligomycin to inhibit phosphorylation. Dark symbols – kinetics of the proton leak subsystem in the absence of an effector (filled circle); open symbols – in the presence of an effector (open circle). Comparing rates of mitochondrial respiration at any given value of membrane potential, one can see that in the presence of the effector the respiratory rate is higher than in its absence so that the whole curve of the dependence of the rate on membrane potential is shifted up. This indicates that proton leak is stimulated by the effector.

3. Calculate the rate of phosphorylation as the difference between the total respiration rate and the rate of proton leak at any given membrane potential value: JP = JC − JL. 4. Plot phosphorylating respiration rate versus Δy as shown in Fig. 6. This represents the kinetics of the phosphorylating subsystem. 5. Repeat these steps with an effector added or with mitochondria affected by some pathological (physiological) agent. 3.6. Determination of Elasticity Coefficients of the Proton Leak, Respiratory, and Phosphorylating Subsystems to Dy

1. Calculate elasticity coefficient of the respiratory subsystem (JC) to Δy using kinetic dependencies of flux through the respiratory subsystem to Δy (Fig. 5). The elasticity of the respiratory subsystem to Δy is determined from the slope of this curve at given value of Δy according to the equation e DCY =

d J C d DY / JC DY

V. Borutaite and R. Baniene

Respiratory rate, nmolO/min mg protein

300 250 200 150 100 50 0 90

100

110 120 130 Membrane potential, mV

140

150

Fig. 5. Kinetics of the respiratory subsystem. Rate of mitochondrial respiration and membrane potential were titrated by carboxyatractyloside. Dark symbols – kinetics of the respiratory subsystem in the absence of an effector (filled circle); open symbols – in the presence of an effector (open circle). Comparing rates of mitochondrial respiration at any given value of membrane potential, one can see that in the presence of the effector the respiratory rate is lower than in its absence so that the whole curve of the dependence of the rate on membrane potential is shifted down. This indicates that the respiratory subsystem is suppressed by the effector. Respiratory rate, nmolO/min mg protein

146

250

200

150

100

50

0 90

95

100

105

110

115

120

Membrane potential, mV Fig. 6. Kinetics of the phosphorylating subsystem. Rate of heart mitochondrial respiration and membrane potential were titrated with rotenone. Dark symbols – kinetics of the phosphorylating subsystem in the absence of an effector (filled circle); open symbols – in the presence of an effector (open symbols). The rate of phosphorylation was estimated as the difference between the total respiration rate and the respiration coupled to proton leak at any given membrane potential value. Comparing rates of mitochondrial respiration at any given value of membrane potential, one can see that in the presence of the effector the respiratory rate is lower than in its absence so that the whole curve of the dependence of the rate on membrane potential is shifted down. This indicates that the phosphorylation subsystem is suppressed by the effector.

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where JC – initial mitochondrial respiratory rate in the state 3 without any inhibitor added. 2. Calculate elasticity coefficient of proton leak (JL) to Δy using kinetic dependence of flux through the proton leak subsystem to Δy (Fig. 4). The slope of this curve at the state 3 membrane potential value (indicated by a dashed line in Fig. 4) is used to calculate the elasticity coefficient of the proton leak to Δy according to the equation: e DLY =

d J L d DY / JL DY

where JL – rate of mitochondrial respiration at membrane potential value corresponding to state 3. 3. Calculate elasticity coefficient of the phosphorylating subsystem (JP) to Δy using kinetic dependencies of flux through phosphorylating system to Δy (Fig. 6). This is determined from the normalized slopes of plots of phosphorylating respiration rate against Δy in the presence of oligomycin (JL) and its absence (JC) during titration of the respiratory chain with rotenone. Use the equation: e DPY = 3.7. Determination of Flux Control Coefficients (C)

d ( J C - J L ) d DY / JP DY

1. Flux control coefficients of the respiratory, the phosphorylating, and the proton leak subsystems over the flux of respiration can be determined using equations: (a) Control coefficient of the respiratory subsystem over respiration: CCJ C =

J P ´ e DPY + J L ´ e DLY J P ´ e DPY + J L ´ e DLY - J C ´ e DCY

(b) Control coefficient of the phosphorylating subsystem over respiration: C PJ C = J P ´

1 - CCJ C JC

(c) Control coefficient of the proton leak subsystem over respiration: J

C LJ C = J L ´

1 - CC C JC

L C P where e DY , e DY , and e DY – elasticities coefficients of proton leak, respiratory, and phosphorylating subsystems to

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Δy; JC – initial mitochondrial respiration rate in the state 3 without any inhibitor added; JP – rate of phosphorylation estimated as the difference in JC − JL at the state 3 membrane potential; JL – rate of proton leak at the state 3 membrane potential. 2. Flux control coefficients of the respiratory, the phosphorylating, and the proton leak subsystems over the phosphorylation flux can be determined using equations: (a) Control coefficient of the respiratory subsystem over of phosphorylation: CCJ P =

J P ´ e DPY

J C ´ e DPY + J L ´ e DLY - J C ´ e DCY

(b) Control coefficient of the phosphorylation subsystem over phosphorylation: J

C PJ P = 1 - J P ´

CC P JC

(c) Control coefficient of the membrane leak subsystem over phosphorylation: J

C LJ P = - J L ´

CC P JC

where e DLY , e DCY , and e DPY – elasticities coefficients of proton leak, respiratory, and phosphorylating subsystems to Δy; JC – initial mitochondrial respiration rate in the state 3 without any inhibitor added; JP – rate of phosphorylation corresponding to the difference in JC − JL at the state 3 membrane potential value; JL – rate of proton leak at the state 3 membrane potential value. Here, we describe how the elasticity and control coefficients of the respiratory, proton leak, and phosphorylating subsystems are estimated at mitochondrial state 3 (maximal mitochondrial respiration rate in the presence of excess ADP). Similarly, the elasticities and control coefficients of the subsystems can be estimated in other states of mitochondrial respiration. Note that control can be distributed differently in various mitochondrial steady states. 3.8. Determination of Flux Control Coefficient of ATP Synthase (CATPsynt )

To determine flux control coefficient of an individual enzyme over a flux through the oxidative phosphorylation system, titrations of respiration with specific inhibitors can be used (“bottom-up” analysis) (see Note 11) (1).

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1. Add 1 mL of the incubation medium to the closed, stirred, and thermostated (37°C) oxygen electrode vessel. 2. Add mitochodria to the vessel (final concentration of mitochondrial protein 0.5 mg/mL). 3. Add 10 μL of 100 mM pyruvate plus 100 mM malate. Monitor mitochondrial respiration for 2–3 min. 4. Then add 10 μL of 100 mM ATP to achieve the state 3 respiration rate (see Note 12). 5. To determine control coefficient of ATP synthase, titrate gradually mitochondrial respiration rate with 0.5–1 μL of oligomycin until oxygen consumption is completely inhibited. 6. Plot a curve of dependence of the respiration rate on the concentration of oligomycin (Fig. 7). 7. Control coefficient of the enzyme can be calculated from the initial slope of the titration curve using equation: C ATPsynthase =

DJ DI J 0 I max

where J0 – initial respiration rate without oligomycin added; ΔJ – difference in respiration rate induced by addition of small amount of oligomycin; Imax – concentration of oligomycin just sufficient to completely inhibit respiration. 250 Respiratory rate, nmolO/min mg protein

9

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50

0

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20

40

Imax

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Oligomycin, nmol/mg protein

Fig. 7. Kinetic dependence of mitochondrial respiration rate on oligomycin concentration. J0 – initial mitochondrial respiration rate in state 3 without any oligomycin added; ΔJ – difference in the respiration rate in the absence and in the presence of 18 nmol/mg oligomycin (ΔI ); Imax – concentration of oligomycin (50 nmol/mg) completely inhibiting respiration.

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4. Notes 1. Nigericin is a electroneutral K+/H+ ionophore; it catalyzes the exchange of one H+ for one K+. An addition of excess of nigericin will clamp ΔpH equal to the K+ gradient, i.e., Δp = Δy. For experiments on liver mitochondria 80 ng/mL nigericin can be used. However, in heart mitochondria nigericin causes inhibition of respiration; therefore, it should be avoided. Do not use nigericin together with valinomycin. Valinomycin induces K+ movement across the membrane, and nigericin will act as a potent uncoupler in the presence of valinomycin dissipating mitochondrial membrane potential (14). 2. Glucose/hexokinase (20 mM glucose and 0–0.6 U/mL hexokinase) can be used as an ADP-regenerating system (8–10). Different rates of ATP turnover and respiration then can be achieved by changing hexokinase concentration. However, one should take into account that at low concentrations hexokinase may become a part of the system exerting significant control. For this reason, creatine/creatine phosphokinase ADPregenerating system is better because creatine phosphokinase is always used in excess. 3. When succinate is used as a respiratory substrate for heart mitochondria 1 μM rotenone should be added. For liver mitochondria, the most preferable substrate is 5 mM glutamate plus 5 mM malate. 4. TPMP+- or TPP+-sensitive electrodes can be manufactured as described in ref. 15. Alternatively, high-resolution respirometer equipped with TPP+-sensitive electrodes can be obtained from Oroboros Instruments (http://www.oroboros.at). TPP+ electrodes are more easy to use as TPP+ equilibrates across the membranes faster than TPMP+; however, TPP+ is more soluble in lipids and may cause more unspecific binding. 5. Before an experiment wash the electrode in distilled water for 0.5–1 h. During calibrations, follow the response of electrode to TPP+. If significant drift or noise of electrode is observed, wash the electrode once again. If drift remains, change the TPP+ selective membrane of the electrode. 6. TPP+ at 1 μM concentrations inhibits respiration of heart mitochondria with NAD-dependent substrates. Mitochondrial respiration with succinate is insensitive to TPP+ up to 8 μM concentration (16). 7. The TPP+ binding correction factor can be determined from the ratio of Rb+ to TPP+ accumulation as a function of mitochondrial volume (15). Alternatively, values reported in literature can be used. For heart mitochondria, the TPP+ binding correction

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factor has been determined to be 0.162 (13) and for liver mitochondria – 0.25 (17). TPMP+ binding correction factor for heart mitochondria was determined to be 0.17 (12) and 0.4 for liver mitochondria (15). 8. When succinate is used as mitochondrial respiratory substrate, respiration rate can be titrated with 0.1–1 mM malonate. 9. When using water-insoluble inhibitors, after each run wash thoroughly the incubation vessel and electrodes with 70% ethanol and then with distilled water. 10. Kinetics of the respiratory subsystem can be measured by changing the creatine/creatine phosphate (e.g., in mM: 5/45, 30/20, 40/10, and 50/0) concentrations in the medium. 11. Specific inhibitors used for determination of flux control coefficients of individual enzymes must be irreversible (1, 4). 12. Check that oxygen consumption rate after ATP addition is linear in the whole range of the oxygen concentrations. If not, change the conditions so that the rate would stay reasonably linear. Lowering the temperature or increasing substrate concentration may help. Alternatively, try adding a single aliquot of the inhibitor when the rate is linear, and repeat this in other incubations with another concentration of inhibitor. References 1. Higgins JJ (1965) Metabolic flux analysis. In: Chance B, Estabrook RW, Williamson JR (eds) Control of energy metabolism. Academic Press, New York, London 2. Kascer H, Burns JA (1973) The control of flux. Symp Soc Exp Biol 27:65–104 3. Fell DA (1992) Metabolic control analysis: a survey of its theoretical and experimental development. Biochem J 286:313–330 4. Brown GC, Lakin-Thomas PL, Brand MD (1990) Control of respiration and oxidative phosphorylation in isolated rat liver cells. Eur J Biochem 192:355–362 5. Hafner RP, Brown GC, Brand MD (1990) Thyroid-hormone control of state-3 respiration in isolated rat liver mitochondria. Biochem J 265:731–734 6. Borutaite V, Mildaziene V, Katiliute Z, Kholodenko B, Toleikis A (1993) The function of ATP/ADP translocator in the regulation of mitochondrial respiration during development of heart ischemic injury. Biochim Biophys Acta 1142:175–180 7. Ciapaite J, Bakker SJ, Van Eikenhorst G, Wagner MJ, Teerlink T, Schalkwijk CG et al (2007) Functioning of oxidative phosphorylation

in liver mitochondria of high-fat diet fed rats. Biochim Biophys Acta 1772:307–316 8. Ciapaite J, Nauciene Z, Baniene R, Wagner MJ, Krab K, Mildaziene V (2009) Modular kinetic analysis reveals differences in Cd2+ and Cu2+ ion-induced impairment of oxidative phosphorylation in liver. FEBS J 276:3656–3668 9. Borutaite V, Morkuniene R, Brown GC (1999) Release of cytochrome c from heart mitochondria is induced by high Ca2+ and peroxynitrite and is responsible for Ca2+-induced inhibition of substrate oxidation. Biochim Biophys Acta 1453:41–48 10. Borutaite V, Morkuniene R, Arandarcikaite O, Jekabsone A, Barauskaite J, Brown GC (2009) Nitric oxide protects the heart from ischemiainduced apoptosis and mitochondrial damage via protein kinase G mediated blockage of permeability transition and cytochrome c release. J Biomed Sci 11:16–70 11. Fell D (1997) Understanding the control of metabolism. Portland Press, London 12. Borutaite V, Mildaziene V, Brown GC, Brand MD (1995) Control and kinetic analysis of ischemia damaged heart mitochondria: which parts of the oxidative phosphorylation system

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are affected by ischemia? Biochim Biophys Acta 1272:154–158 13. Mildaziene V, Baniene R, Nauciene Z, Marcinkeviciute A, Morkuniene R, Borutaite V et al (1996) Ca2+ stimulates both the respiratory and phosphorylation subsystems in rat heart mitochondria. Biochem J 320:329–334 14. Brown GC, Brand MD (1986) Changes in permeability to proton motive force in rat liver mitochondria. Biochem J 234:75–81 15. Brand MD (1995) Measurement of mitochondrial protonmotive force. In: Brown GC,

Cooper CE (eds) Bioenergetics: a practical approach. Oxford University Press, New York 16. Mildaziene V, Baniene R, Marcinkeviciute A, Nauciene Z, Kalvenas A (1997) Tetraphenylphosphonium inhibits oxidation of physiological substrates in heart mitochondria. Mol Cell Biochem 174:67–70 17. Hoek JB, Nicholls DG, Williamson JR (1980) Determination of the mitochondrial protonmotive force in isolated hepatocytes. J Biol Chem 255:1458–1464

Chapter 10 Expression of Uncoupling Proteins in a Mammalian Cell Culture System (HEK293) and Assessment of Their Protein Function Martin Jastroch Abstract Immortalised cultured cells are powerful tools to assess the function of ectopically expressed proteins. However, it must be ensured that the protein of interest is functional in the host system and display native behaviour. In particular, mitochondrial uncoupling proteins (UCPs) displayed (non-native) artefactual uncoupling when expressed in yeast or can possess functions upon reconstitution in proteoliposomes that cannot be reproduced in isolated mitochondria. In the light of newly discovered UCP1 orthologues and paralogues (UCP2, UCP3, plant UCP), comparative functional studies require a system with identical mitochondrial, cellular, and genetic backgrounds. In this chapter, the protocols for the ectopic expression of mouse UCP1 in the human embryonic kidney (HEK293) cell line are introduced. In isolated cell culture mitochondria, the proton leak can be measured and modulators of UCP1 activity can be tested. As mouse UCP1 in this system shows native behaviour, this may be a suitable system to directly compare the functional relationships between different UCPs. Key words: Human embryonic kidney cells, Uncoupling proteins, Proton leak kinetics, Proton leak

1. Introduction Uncoupling protein 1 (UCP1) is a 32-kDa protein belonging to the superfamily of mitochondrial anion carrier proteins residing in the mitochondrial inner membrane. UCP1 catalyses a net proton leak and thereby dissipates mitochondrial proton-motive force as heat in brown adipose tissue (1). Its activity is inhibited by physiological concentrations of purine nucleoside di- and triphosphates and inhibition can be overcome by free fatty acids. Although

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_10, © Springer Science+Business Media, LLC 2012

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UCP1 is the best understood member of the uncoupling protein family, it remains unclear how fatty acids contribute to net proton transport and whether the protein directly translocates protons. Currently, there are three competing models to explain UCP1 function (2–5). UCP1 is the archetypal uncoupling protein and thought to be expressed exclusively in mammalian brown adipose tissue. Based on sequence identity and conserved genomic synteny, other UCPs in different tissues and vertebrate species were annotated as UCP1 orthologues (6) and paralogues (UCP2, UCP3, and plant UCP) (7–9). There is currently no agreement on their protein function and physiological role. A direct comparison of their molecular mechanisms, activity, and regulation would greatly assist in annotating their protein function. This is, however, impossible in native tissues due to differences in tissue specificity, differential abundance, and presence in different animal species. UCP-ablated mice resemble a system where different phenotypes can be studied, but confounding differences in the life history and bioenergetic status in response to UCP-ablation may also result in distracting, secondary cellular adaptations. A number of molecular experimental systems have been applied in the past to identify and compare the function of different UCPs. Proteoliposomes with reconstituted UCPs were used to identify transported metabolites, sequence motifs, and modulators of UCP function. However, the results obtained from proteoliposomes have to be further substantiated in other systems, as the physiological significance for some observations had to be rejected. For example, coenzyme Q was identified as an obligatory cofactor of UCP1 activity in proteoliposomes (10), but is not required in yeast mitochondria containing UCP1 (11). Furthermore, UCP3 shows GDP-inhibition of proton translocation in proteoliposomes (12), which is absent in skeletal muscle mitochondria (13). In the yeast cell system, UCP1 appears to function natively at low levels while high protein concentration results in artefactual, GDP-insensitive uncoupling (14). But even in yeast strains with moderate UCP1 expression, substrate oxidation appears to be depressed by UCP1 expression and some proportion of uncoupling is not GDP-sensitive when compared to wild-type strains (15). Here, a new cell system is introduced, expressing UCPs in human embryonic kidney (HEK293) cell culture. It is exemplified how to generate a stable cell line that ectopically expresses mouse UCP1, how to isolate mitochondria from tissue cell culture, and how to measure UCP1 protein function. Mouse UCP1 demonstrably displays native behaviour (fatty acid and GDP sensitivity) without artefactual uncoupling (16).

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2. Materials 2.1. Cell Culture Conditions for HEK293 Cells

1. Cell culture medium: Dulbecco’s modified eagle medium (DMEM; GIBCO), 4,500 mg/L glucose, +L-glutamine, -pyruvate, supplemented with 10% FCS (BIOCHROM), 50 mg/mL gentamycin (GIBCO) and 2.5 mg/mL amphotericin B (SigmaAldrich). 2. Cryo-medium: Cell culture medium supplemented with 10% DMSO. 3. Trypsin (0.2%) dissolved in PBS pH 7.5.

2.2. Transfection and Selection for Stably Transfected Cells

1. Superfect® Transfection Reagent (QIAGEN). 2. Geneticin (GIBCO) stocks are dissolved in ultrapure water at 400 mg. 3. Silicon tubing (diameter: 5–10 mm) is evenly cut to small rings of 3–5 mm length and soaked in ethanol for sterilisation (and washed in cell culture medium before use).

2.3. Sample Preparation, SDS-PAGE, Western Blotting, and Immunological Detection

1. 2× concentrated sample buffer (0.125 M Tris–Cl at pH 6.8, 4% w/v SDS, 0.2 M DTT, 20% v/v glycerol, 0.2% w/v bromophenol blue), stored at −20°C. RIPA buffer (50 mM Tris–Cl, 1% NP-40, 0.25% sodium-deoxycholate, 150 mM sodium chloride, 1 mM EDTA, 1 mM sodium-orthovanadate, 1 mM phenylmethylsulfonyl fluoride, stored at 4°C, Protease inhibitor Cocktail III (CALBIOCHEM), added on the experimental day). 2. Gel electrophoresis: 8× separation buffer (3 M Tris–Cl pH 8.8), 8× stacking buffer (0.5 M Tris–Cl pH 6.8), 10× running buffer (0.25 M Tris-Glycine pH 8.3, 1% SDS). 3. Blotting: Transfer buffer (48 mM Tris-Glycine, 1.3 mM SDS, adjusted to pH 9.2). 4. Staining of the Western membrane: Ponceau S staining solution (0.2% Ponceau S (w/v), 3% (v/v) trichloric acid). 5. Immunological detection: 10× TBS (0.2 M Tris, 80 g/L sodium chloride at pH 7.6), blocking solution (1× TBS, 0.1% Tween, 5% milk powder) (see Note 1), washing solution (1× TBS, 0.1% Tween), ECL-Kit (Amersham).

2.4. Mitochondrial Isolation

1. Isolation buffer (0.25 M sucrose, 5 mM Tris–Cl, 2 mM EGTA, adjusted to pH 7.4 at 4°C). When required, 0.5% defatted BSA (A3803, SIGMA) is added and dissolved in the buffer. 2. Gauze or nylon netting for filtering, soft hairbrush for resuspension of pellets. 3. Cooling centrifuge with a rotor of minimum 11,000 × g capacity, 40 mL centrifuge tubes.

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2.5. Measurement of Mitochondrial Oxygen Consumption and Proton Leak

1. The experiments are performed in a set-up obtained from Rank Brothers Ltd, Cambridge, UK. A 7.0-mL reaction chamber is connected to a circulating water bath maintained at 37°C. A TPMP+ sensitive electrode and a lid for the TPMP+ sensitive electrode are custom-made as described in Chapter 11. The TPMP+ electrode and a reference electrode are connected to a pH amplifier. Both oxygen and TPMP+ signal are digitalised using a PowerLab (ADinstruments). 2. Assay buffer (120 mM KCl, 3 mM HEPES, 5 mM KH2PO4, 1 mM EGTA adjusted to pH 7.2, 0.3% of defatted BSA is dissolved at the experimental day). 3. Inhibitors and drugs: 4 mM rotenone, 1 mg/mL oligomycin, 100 mM nigericin, 1 mM FCCP, all dissolved in EtOH. 4. Substrates and TPMP+: 1 M succinate, 0.5 M malonate, dissolved in assay buffer at pH 7. A 10 mM and a 0.5 mM TPMP+ stock dissolved in water.

3. Methods The coding sequence of mouse UCP1 was transcribed from mRNA to cDNA using reverse transcriptase and cloned into an expression vector suitable for stable transfection of mammalian cell culture (pcDNA3.1, containing neomycin/geneticin resistance). A subconfluent HEK293 cell culture dish was transfected with either an empty vector or a vector containing mouse UCP1 and then selected for stable transfection. Clonal cells from single colonies were picked and grown separately on 12-well plates (for detail, see Subheading 3.1). When confluent, the cells were harvested and transferred to two 10-cm cell culture dishes. The cells of one confluent 10-cm dish were collected and stored in liquid nitrogen, while the other dish was frozen at −80°C for Western blot analysis, testing for positive transfection (see Subheading 3.2, Fig. 1a). Positive cells were thawed and further grown on 10 cm dishes for maintenance. When confluent, a 10-cm dish was harvested and cells were stored in four 1 mL aliquots in liquid nitrogen (see Subheading 3.3). For the assessment of UCP function, a vial of empty vector and mouse UCP1-containing vector was thawed and cells were grown (see Subheading 3.4) to a 2,000 cm2 monolayer before mitochondria were isolated by differential centrifugation (see Subheading 3.5). The protein content of the mitochondrial pellet was determined by the biuret method. A sample was added to a temperaturecontrolled chamber with a Clark-type electrode to measure respiratory control ensuring mitochondrial integrity after the isolation process (respiratory control ratio is typically around 3). For the measurement of proton leak kinetics (see Subheading 3.6), 0.35 mg of mitochondria was added and the commercial lid of the reaction

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Fig. 1. Autoradiography of Western blots probed for mouse UCP1. (a) Fifty micrograms of cell homogenate were blotted and probed with a rabbit anti-hamster UCP1 antibody. A cell line with stably transfected mouse UCP1 (+) is compared to a wild-type cell line (−). (b) Ten micrograms of UCP1-containing HEK cell mitochondria are blotted and UCP1 detected. The right-hand lane (“Cell debris”) was loaded with cell debris that was collected from the pellet of the slow centrifugation step (after second homogenisation). (b) illustrates that UCP1 is incorporated in the mitochondrial membrane and not in other cell compartments or membranes. MM molecular marker (Magic mark).

chamber was replaced with a lid enclosing a TPMP+ sensitive and a reference electrode (see Chapter 11). The simultaneous measurement of oxygen consumption and membrane potential allowed the calculation of the proton leak. To convert the oxygen consumption into proton leak rate, the mitochondria were measured in the presence of oligomycin and energised with succinate. Oligomycin inhibited the ATP-synthase and the residual respiration can be solely addressed to leak. When mitochondria are energised with succinate, it is assumed that six protons per atomic oxygen are transported from the matrix to the intermembrane space (and leak back). To measure the kinetics of the leak, substrate oxidation was sequentially inhibited with malonate (competitively inhibiting the succinate dehydrogenase) to record the steady states of oxygen consumption and membrane potential simultaneously. Plotting oxygen consumption against membrane potential illustrates the proton leak kinetics. At a common membrane potential, two proton leak rates could be directly compared. Identical measurements were performed in UCP1-containing and wild-type mitochondria in the absence and presence of guanosine diphosphate to address the contribution of UCP1 to mitochondrial proton conductance (Fig. 2). The residual mitochondria are subjected to Western blot analysis to confirm UCP1 expression and protein content (Fig. 1b) 3.1. Transfection of HEK293 Cells

1. HEK293 cells are stored in 1–1.5 mL aliquots of cryo-medium in liquid nitrogen. An aliquot is quickly thawed by incubation in a 37°C water bath and seeded on a sterile 10-cm cell culture dish with 10 mL culture medium. The medium is replaced after 1 h to allow better recovery of the HEK293 cells.

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Fig. 2. Proton leak kinetics of wild-type and UCP1-containing HEK293 mitochondria. The proton leak curves were deduced from original data [16] by best-fitting of the data points to an exponential function. The upward shift of the red curve (UCP1-containing HEK mitochondria) demonstrates an increased leak. At a common membrane potential, the oxygen consumption rate (as a measure for proton leak rate) of UCP1-containing mitochondria (red curve) is higher as in wild-type HEK mitochondria (yellow curve). Importantly, when UCP1-containing mitochondria were measured in the presence of GDP (0.5–1.0 mM, green curve), the UCP1 curve overlaid with the wild-type curve. This demonstrates that UCP1 could be completely inhibited in the HEK293 cell system. No difference in basal proton conductance between UCP1 containing and wild-type HEK293 cells excludes artefactual uncoupling by mouse UCP1. This suggests that UCP1 is properly folded and correctly incorporated into the mitochondrial inner membrane.

2. The cells are passaged when the cell culture dish has grown confluent. The medium is removed, the cells washed once with PBS and 2 mL of trypsin buffer is added. When the cells detach from the bottom of the plate, the trypsin reaction is stopped by adding excess cell culture medium. The cells from one plate can be seeded to five to eight new dishes. After seeding, the cell culture medium is replaced 1 h later (see Note 2). 3. At 50–70% confluency, the cells are transfected with the plasmid. Before adding the transfection mix, the cells are washed in PBS a few times. 4. The plasmid (2–5 mg) is diluted with DMEM (without serum) to a final volume of 150 ml, and 4 ml/mg plasmid of superfect (QIAGEN) is added. The reaction is mixed vigorously, incubated for 10 min and 2 mL of complete cell culture medium is subsequently added. This reaction mix is pipetted to the washed cells so that it just covers all cells (about 2.4 mL) and the plate is incubated at 37°C. After 3 h, the cells are rinsed with PBS three times and the dish is filled up with 10 mL of culture medium.

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5. After 24 h of recovery, the cell culture medium is replaced with fresh medium containing 400 mg/mL geneticin (GIBCO). In parallel, a cell culture plate with non-transfected cells is treated similarly with geneticin and serves as a selection control. 6. During the selection process, the geneticin-containing cell culture medium is replaced every 3–4 days. This process continues until all cells on the non-transfected control plate die and single colonies can be spotted on the transfected plates. 7. When the colonies have grown to a diameter of 1–2 mm, the cell culture medium is removed and the plate is washed two times with PBS. A small silicon ring is pressed and held down to the bottom of the cell plate around the colony with sterile tweezers and filled with trypsin containing buffer. After 1 min, the solution is pipetted up and down to detach all cells of the colony and transferred in reaction tubes containing 1 mL warm cell culture medium. This process is repeated for a maximum number of colonies. 8. The cells of single colonies are transferred to a 12-well plate and regrown to confluency in the presence of geneticin, then transferred to two 10-cm culture dishes. 9. Once confluent, cells of one dish are washed, trypsinised, and slowly frozen in cryo-medium (see Note 3), while the other dish was directly frozen at −80°C for immunological detection of UCP1. 3.2. Sample Preparation, SDS-Page, Western Blotting, and Immunological Detection of UCP1

1. For the detection of UCP1 from whole cell lysates, the frozen cells are scraped (see Note 4) from the plates with 1 mL of icecold RIPA buffer and filled into a 1.5 mL reaction tube. The cells are then shaken for 15 min at 4°C to lyse the cells. Then, the reaction is centrifuged at 14,000 × g at 4°C for 15 min and the protein-containing supernatant is transferred to a new reaction tube for further analysis. Fifty micrograms of whole cell protein is sufficient for the detection of UCP1. One volume of 2× sample buffer is added, incubated at 99°C for 5 min (see Note 5) and cooled rapidly on ice. 2. For the detection of UCP1 from mitochondrial protein, 5–10 mg of mitochondrial protein are mixed with the equal volume of 2× sample buffer, incubated at 99°C for 5 min and cooled rapidly on ice. 3. For gel electrophoresis, a vertical chamber system (BIORAD) was used. A 10% separation gel is prepared by mixing 3.33 ml 37.5:1 acrylamide/bis- (ROTH), 1.25 mL 8× separation buffer, 5.27 mL double-distilled water, 0.1 mL 10% SDS, 50 mL 10% ammonium persulfate (ROTH), 5 mL N,N,N,N¢tetramethyl-ethylenediamine (TEMED, ROTH). After polymerisation, a 3% stacking gel is prepared by mixing 0.5 mL

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37.5:1 acrylamide/bis-, 1.25 mL 8× separation buffer, 3.16 mL double-distilled water, 50 mL 10% SDS, 30 mL 10% ammonium persulfate, 10 mL N,N,N,N¢-tetramethyl-ethylenediamine (TEMED, ROTH). 4. After polymerisation, the gel is placed in running buffer and the sample as well as a marker (here: Magic Mark, INVITROGEN) are loaded into the gel pockets. The gel can be run until the bromophenol blue band leaves the lower end of the gel. 5. The gel is transferred via a semi-dry blotting system (Transblot SD, BIORAD) onto a nitrocellulose membrane (Hybond ECL, AMERSHAM). The gel, membrane, and filterpaper used during blotting have been soaked in transfer buffer. 6. An electrical current of 0.01 mA/cm2 is applied for 30 min to transfer the proteins from the gel to the membrane. 7. To control for protein transfer, the membrane is stained with Ponceau S for 5 min, briefly washed in double-distilled water and scanned for densitometry (see Note 6) and documentation. 8. For detection of UCP1, the membrane is first incubated in blocking solution for an hour at room temperature or alternatively, overnight at 4°C to reduce non-specific binding of the antibody. All incubation and washing steps are carried out while gently shaking. Then, the membrane is washed briefly two to three times in washing solution. An anti-UCP1 antibody (in our lab a rabbit-anti-hamster UCP1 antibody, see also Note 7) is diluted 1:10,000 in washing buffer and the membrane incubated for an hour at room temperature or overnight at 4°C. 9. The membrane is then incubated once for 15 min, three times for 5 min in washing solution at room temperature. 10. A secondary antibody (in our lab: a goat-anti-rabbit, HRPconjugated) is added at a 1:10,000 dilution in washing solution and washing steps repeated as for the primary antibody. 11. To detect the antibody signal, the membrane is incubated for 5 min in enhanced chemoluminiscence (ECL) solution (AMERSHAM) and then wrapped in transparent covering and exposed to an X-ray film. 3.3. Preparation of HEK Cell Cryo-Stocks

The cell culture medium is aspirated and the cells are washed twice with PBS, 1 mL of trypsin buffer is then added to detach the cells and the reaction is stopped with 3 mL of cell culture medium. 900 mL of cells is then added to a cell culture tube with 100 mL of DMSO and the tube gently inverted. For the freezing procedure, the cells are slowly cooled down to −80°C (see Note 3) and then transferred to liquid nitrogen for long-term storage.

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To grow cells for the functional assay, a vial of HEK293 cells is thawed as described in Subheading 3.1, step 1. Upon confluency, cells were passaged to three 10 cm plates (after ~4th day) and those passaged to eight 10-cm plates (after ~7th day). Cells from the eight 10 cm plates were harvested (after ~9th day) and seeded onto four 500 cm2 plates (CORNING). The cell medium was changed after 3 days and the cells collected 1 day later for mitochondrial isolation. 1. The 500-cm2 cell culture plate is carefully washed twice with ice-cold PBS. All steps from here on are carried out at 4°C. Approximately 10 ml of PBS is pipetted to the plate; cells are collected with a cell culture scraper and filled into a 40-mL centrifuge tube. This procedure is repeated for all plates. 2. The cell suspension is centrifuged at 500 × g for 5 min, the supernatant is discarded and the loose pellet placed in a glass– glass homogeniser (Dounce). 3. The homogeniser is filled with isolation medium containing 0.5% defatted BSA. 4. The cells are homogenised with ten strokes (see Note 8), and the homogenate is refilled into the centrifuge tube and centrifuged at 1,000 × g for 10 min. 5. The supernatant is filtered through gauze or a nylon net (diameter >0.4 mM) into the next centrifuge tube without disturbing the pellet, and then centrifuged at 10,500 × g for 10 min. Meanwhile, the residual pellet is homogenised again by repeating steps 4 and 5. 6. The centrifuge tube from the fast-spin should contain a brownish mitochondrial pellet. The supernatant is discarded and the pellet carefully re-suspended in isolation medium using a soft hairbrush. 7. The homogenate is centrifuged again at 10,500 × g for 10 min, in parallel with the second homogenate. 8. After this fast-spin, step 6 is repeated and both re-suspended pellets are combined. The combined homogenate is centrifuged at 10,500 × g for 10 min and the pellet re-suspended in a minimum volume of isolation buffer. 9. The homogenate is now ready for protein quantification. Using the Biuret method, a dense homogenate typically contains 10 mg/mL mitochondrial protein, with a total yield of 4–8 mg protein. The mitochondrial suspension is kept on ice.

3.6. Respiratory Control Ratio and Proton Leak Kinetics

1. For the measurement of mitochondrial respiration and respiratory control ratios (RCRs), 0.3 mg of mitochondria was added to 2 mL of assay buffer in temperature controlled chamber containing a Clark-like electrode embedded at the bottom. A tight lid ensures an almost closed system. A final concentration

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of 5 mM rotenone is added to prevent uncontrolled oxidation of NADH-linked substrates. 4 mM succinate is added to start the reaction. When respiration stabilises, ADP (600 mM) is added to induce ATP synthesis and increase respiration. When the oxygen consumption rate is stable, 1 mg/mL oligomycin is added to inhibit the ATP synthase and the respiration observed is due to the mitochondrial proton leak. The ratio of ATP synthesising respiration and leak respiration is typically around 3 for wild-type HEK293 mitochondria and reports the integrity of the mitochondria after the isolation process. 2. For the measurement of mitochondrial proton leak, a set-up as described in Chapter 11 is used. Mitochondria (0.7 mg) are added to 2 mL of assay buffer in the reaction chamber. Subsequently, 5 mM rotenone (to prevent oxidation of NADH-linked substrates), 100 nM nigericin (to abolish the pH gradient), and 1 mg/mL oligomycin (to inhibit the ATP synthase) are added. Then, a lid including the TPMP+ and a reference electrode is fitted on top of the chamber to drive out residual water bubbles. 3. The TPMP electrode is calibrated by sequential steps of 0.5 mM TPMP+ up to 2.5 mM. The reaction is started by addition of 4 mM succinate. As soon as mitochondrial respiration and membrane potential reach steady state conditions, multiple additions of malonate (typically 5, up to 2 mM) are made to sequentially inhibit respiration and measure the oxygen consumption rate (which reports the proton leak reate) at different membrane potentials. At the end of the measurement, 0.3 mM FCCP is added to release TPMP+ from the mitochondria and correct for electrode drift. 4. The analysis and illustration of the proton leak kinetics is described in Chapter 11. 5. To measure the effect of UCP1 on proton leak per se, as well as the effect of mall molecules on UCP1 (in this case: guanosine diphosphate), the measurements are conducted in mitochondria ± UCP1. The small molecules are added at the beginning of the measurements, after addition of rotenone, nigericin, and oligomycin. Figure 2 shows proton leak curves derived from original data points and the effect of GDP.

4. Notes 1. Instead of milk powder, Slimfast® appears to block unspecific signals of the UCP1 antibody better and reduces cross-reactions during UCP detection 2. If the cells are attached after passaging, they should be checked with a light microscope.

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3. To achieve better cell viability, the cells should be cooled down slowly to −80°C using a cooling rack (e.g. Mr. Frosty, NALGENE). Thawing of the cells, however, must occur rapidly by placing the cryotube in a 37°C water bath. 4. Collection of the cells is best done using a police rubber-man, which is also called cell scraper. 5. The results of the protein detection are usually of better quality when the reaction is shaked vigorously during incubation using a thermo-shaker. 6. The quantified Ponceau stain can be used as a loading control. 7. A commercial antibody to detect mouse UCP1 is available from Sigma-Aldrich. In some of our studies, we successfully used a rabbit anti-UCP1 polyclonal antibody (1:30,000) from CHEMICON (Cat. No. 3046). 8. If the respiratory control ratios of the mitochondria are low, they are most likely damaged during the mitochondrial isolation. The most critical step of the isolation is the homogenisation step. In case of damage, the number of strokes may be reduced or the pestle of the Dounce homogeniser can be replaced with a pestle of greater clearance.

Acknowledgements The author would like to thank Dr. Sung Won Choi for advice to improve transfection protocols and Ajit Divakaruni for helpful discussion. M.J. is supported by the Deutsche Forschungsgemeinschaft (JA 1884/2-1). References 1. Locke RM, Nicholls DG (1984) Thermogenic mechanisms in brown fat. Physiol Rev 64: 1–64 2. Klingenberg M, Winkler E (1985) The reconstituted isolated uncoupling protein is a membrane potential driven H+ translocator. EMBO J 4:3087–3092 3. Garlid KD, Orosz DE, Modrianský M, Vassanelli S, Jezek P (1996) On the mechanism of fatty acid-induced proton transport by mitochondrial uncoupling protein. J Biol Chem 271:2615–2620 4. Rial E, Aguirregoitia E, Jiménez-Jiménez J, Ledesma A (2004) Alkylsulfonates activate the uncoupling protein UCP1: implications for the transport mechanism. Biochim Biophys Acta 1608:122–130

5. Shabalina I, Jacobsson A, Cannon B, Nedergaard J (2004) Native UCP1 displays simple competitive kinetics between the regulators purine nucleotides and fatty acids. J Biol Chem 279:38236–38248 6. Jastroch M, Withers KW, Taudien S, Frappell PB, Helwig M, Fromme T et al (2008) Marsupial uncoupling protein 1 sheds light on the evolution of mammalian nonshivering thermogenesis. Physiol Genomics 32:161–169 7. Fleury C, Neverova M, Collins S, Raimbault S, Champigny O, Levi-Meyrueis C et al (1997) Uncoupling protein-2: a novel gene linked to obesity and hyperinsulinemia. Nat Genet 15:269–272 8. Boss O, Samec S, Paoloni-Giacobino A, Rossier C, Dulloo A, Seydoux J et al (1997) Uncoupling

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protein-3: a new member of the mitochondrial carrier family with tissue-specific expression. FEBS Lett 408:39–42 9. Laloi M, Klein M, Riesmeier JW, Müller-Röber B, Fleury C, Bouillaud F et al (1997) A plant cold-induced uncoupling protein. Nature 389:135–136 10. Echtay KS, Winkler E, Klingenberg M (2000) Coenzyme Q is an obligatory cofactor for uncoupling protein function. Nature 408:609–613 11. Esteves TC, Echtay KS, Jonassen T, Clarke CF, Brand MD (2004) Ubiquinone is not required for proton conductance by uncoupling protein 1 in yeast mitochondria. Biochem J 379: 309–315 12. Echtay KS, Winkler E, Frischmuth K, Klingenberg M (2001) Uncoupling proteins 2 and 3 are highly active H(+) transporters and highly nucleotide sensitive when activated by coenzyme Q (ubiquinone). Proc Natl Acad Sci USA 98:1416–1421

13. Parker N, Affourtit C, Vidal-Puig A, Brand MD (2008) Energization-dependent endogenous activation of proton conductance in skeletal muscle mitochondria. Biochem J 412: 131–139 14. Stuart JA, Harper JA, Brindle KM, Jekabsons MB, Brand MD (2001) A mitochondrial uncoupling artifact can be caused by expression of uncoupling protein 1 in yeast. Biochem J 356:779–789 15. Esteves TC, Parker N, Brand MD (2006) Synergy of fatty acid and reactive alkenal activation of proton conductance through uncoupling protein 1 in mitochondria. Biochem J 395:619–628 16. Jastroch M, Hirschberg V, Brand MD, Liebig M, Weber K, Bolze F et al (2006) Introducing a mammalian cell system to study the function of evolutionary distant uncoupling proteins. Biochim Biophys Acta 371–372

Chapter 11 Measurement of Proton Leak and Electron Leak in Isolated Mitochondria Charles Affourtit, Casey L. Quinlan, and Martin D. Brand Abstract Oxidative phosphorylation is an important energy-conserving mechanism coupling mitochondrial electron transfer to ATP synthesis. Coupling between respiration and phosphorylation is not fully efficient due to proton and electron leaks. In this chapter, methods are presented to measure proton and electron leak activities in isolated mitochondria. The relative strength of a modular kinetic approach to probe oxidative phosphorylation is emphasised. Key words: Mitochondria, Oxygen consumption, Membrane potential, Reactive oxygen species production, Oxidative phosphorylation, Modular kinetic analysis, Proton leak, Electron leak

1. Introduction Mitochondria conserve energy as ATP by oxidative phosphorylation. Reducing equivalents derived from the cellular breakdown of carbonbased substrates are donated to the mitochondrial electron transfer chain and eventually fully reduce molecular oxygen to water. The energy liberated during this mitochondrial electron transfer is used to establish an electrochemical proton gradient across the mitochondrial inner membrane that in turn is used to drive ATP synthesis (1). The coupling between electron transfer and ATP synthesis is not absolute: protons can flow back into the mitochondrial matrix by mechanisms that bypass the ATP synthase (2) and electrons derived from respiratory complexes I and III (predominantly) can be used to incompletely reduce oxygen to superoxide (3). Although these proton and electron leaks lower the coupling efficiency of oxidative phosphorylation and thereby decrease the ATP that is conserved from carbon fuels, they are not necessarily wasteful processes. Modulation of coupling efficiency by proton leak Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_11, © Springer Science+Business Media, LLC 2012

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activity is a likely and important mechanism to control cell physiology, which is for example evident in pancreatic beta cells where glucose-stimulated insulin secretion is regulated by uncoupling protein 2, a significant contributor to beta cell proton leak (4). Mitochondria-derived superoxide and other reactive oxygen species are key signals in cellular redox regulation, and their production is, therefore, also highly relevant in terms of cell physiology (5). It is clear that mitochondrial proton leak and electron leak are important processes and that they, therefore, require accurate and reliable measurement. In this chapter, we present protocols to determine proton and electron leak activities in rat skeletal muscle mitochondria. Moreover, we describe a procedure to obtain a system-level, modular kinetic description of oxidative phosphorylation. All methods may be readily adapted for use with mitochondrial systems other than rat skeletal muscle, but some caution is warranted. The presented Amplex Red assay, for example, only reports superoxide production rates in experimental systems that exhibit endogenous ROS scavenging activity that is sufficiently low for the added hydrogen peroxidase to compete (this, for example, excludes crude rat liver preparations from this assay).

2. Materials Unless stated otherwise, all chemicals may be purchased from Sigma-Aldrich. 2.1. Mitochondrial Isolation from Rat Skeletal Muscle

1. Animals – Wistar rats, aged between 5 and 8 weeks. 2. Equipment – Sharp scissors and razor blades; cutting board, Polytron homogenizer; ten 50-mL Falcon tubes; eight 50-mL centrifuge tubes; two square pieces of muslin or cheese cloth; full-size refrigerated centrifuge. 3. Solutions – Chappell-Perry buffer 1 (CP1): 100 mM KCl, 50 mM Tris–HCl (pH 7.2 at room temperature) and 2 mM EGTA; Chappell-Perry buffer 2 (CP2): 100 mM KCl, 50 mM Tris–HCl (pH 7.4 at 4°C), 2 mM EGTA, 0.5% BSA, 5 mM MgCl2, 1 mM ATP, and 250 units/100 mL Protease Type VIII. 4. Protein assay – Biuret reagent or similar protein measurement system for determination of mitochondrial yield and concentration.

2.2. TPMP-Selective Electrode Sleeve

1. Solutions – Tetraphenylboron (10 mM) in 6-mL tetrahydrofuran (THF); 1 g high-molecular-weight polyvinylchloride (PVC) in 20 mL THF; dioctylphthalate (a plasticiser); TPMP (10 mM in H2O stored at room temperature). 2. Equipment – Five glass Petri dishes (10 cm diameter); PVC tubing (4 mm outside diameter); sharp razor blades and scissors.

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1. Oxygen consumption – A water-jacketed Clark oxygen electrode, oxygen-permeable Teflon membrane (see Note 1), an electrode-controller unit and an electronic stirrer that may all be obtained from Rank Brothers Ltd (Cambridge, UK); a circulating water bath. 2. Membrane potential (Δψ) – A TPMP (methyltriphenylphosphonium)-selective electrode sleeve (cf. Subheadings 2.2 and 3.2); a 2–3-cm piece of platinum wire soldered to screened cable; a solid-state Ag/AgCl reference electrode (World Precision Instruments Inc., USA); an adapted oxygen electrode plunger with two additional holes that allow insertion of the TPMP+ and Ag/AgCl reference electrodes; a pH meter (we use a pH-Amp front-end from ADInstruments, UK). 3. Data acquisition – A digital recording system comprised of a PowerLab analog-to-digital signal converter (ADInstruments, UK) linked to any personal computer running LabChart software (ADInstruments, UK). 4. Assay buffer – KHEP (115 mM KCl, 10 mM KH2PO4, 3 mM HEPES, pH 7.2, 1 mM EGTA, 2 mM MgCl2, 0.3% w/v bovine serum albumin) stored at 4°C (see Note 2). 5. Calibration – Sodium dithionite (solid) and TPMP (1 mM in H2O), both kept at room temperature, to calibrate the oxygen and TPMP-selective electrodes, respectively. 6. Respiratory substrates and effectors – Succinate, pyruvate, malate, malonate (free acid stocks at 1 M), ADP (100 mM), all prepared in KHEP with pH adjusted to 7.2. All stocks are stored at −20°C 7. Effectors – 1 mg/mL Nigericin, 4 mM rotenone, 1 mg/mL oligomycin and 1 mM carbonylcyanide p-trifluoromethoxyphenylhydrazone (FCCP), all prepared in ethanol (96%). All stocks are stored at −20°C. 8. Assays – Automatic pipettes to add KHEP and mitochondria; Hamilton microsyringes (5–25 μL) to add respiratory substrates and effectors.

2.4. Mitochondrial Superoxide Production

1. Equipment – A computer-driven and temperature-controlled spectrofluorometer that is capable of acquiring, processing, and storing data, scanning a wavelength range of 220–900 nm and that has excitation, emission, and synchronous spectrum as well as time-course measurement capabilities (Shimadzu and Varian make suitable models); electronic stirrer and stir bars; Hamilton syringes; quartz or disposable cuvettes. 2. H2O2 detection – Amplex Red (Invitrogen) (10 mM in DMSO), horseradish peroxidase (1,000 U/mL in KHE), superoxide dismutase (5,000 U/mL in KHE).

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3. Calibration – H2O2 stock solution (1 mM in H2O; see Note 3). 4. Assay buffer – KHE (115 mM KCl, 3 mM HEPES, pH 7.2, 1 mM EGTA, 0.3% w/v BSA), stored at 4°C and pre-heated to 37°C before use. 5. Substrate and inhibitor stock solutions – 1 M Succinate, 1 M malate, 1 M glutamate prepared in KHE; 4 mM rotenone, 2.5 mM antimycin A and 0.5 mM stigmatellin prepared in ethanol (96%). All stocks are stored at −20°C.

3. Methods In this chapter, we provide methods to measure proton leak during oxidative phosphorylation in isolated rat skeletal muscle mitochondria. We also describe assays to assess the possible electron leak that may occur at respiratory complexes I and III. No protocols are given to probe the precise molecular mechanisms of these proton and electron leaks, which are indeed incompletely understood at present. The standard operating procedure in our laboratory is to determine generic proton leak activity at a range of Δψ values. This procedure yields proton leak kinetics that are an integral part of a more complete, system-level kinetic description of oxidative phosphorylation. In Subheading 3.5, we describe an approach to arrive empirically at such a system-kinetic picture. 3.1. Mitochondrial Isolation from Rat Skeletal Muscle

The following protocol is based on methods described by Ashour et al. (6) and Letellier et al. (7). 1. Anaesthetise one or two rats by approximately 1–2 min exposure to CO2 gas in a closed chamber. Sacrifice the animal(s) by cervical dislocation. 2. Dissect out skeletal muscle by cutting around hind limbs with sharp scissors and peeling back skin from the fascia layer. Quickly retrieve as much muscle as possible, including some from the animal’s dorsal area, and put it immediately in chilled CP1 medium kept on ice. 3. Discard the CP1 medium after dissection is completed and weigh the tissue; yields are typically 15–25 g/rat. After weighing, return the tissue immediately to fresh chilled CP1 medium kept on ice. 4. Take one piece of tissue (or more if the tissue was dissected into small pieces) and place on a pre-cooled cutting board. Trim fat and connective tissue away from the muscle. Finely chop the remaining tissue with sharp scissors and mince it thoroughly with new razor blades till a homogeneous mass without visible lumps is obtained. Place the minced tissue in fresh CP1 medium kept on ice.

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5. Repeat step 4 to mince all tissue. 6. Mix the minced tissue in CP1 medium and allow it to settle. Then, carefully decant the medium thereby removing floating pieces of connective tissue. Rinse the tissue 4–5 times in this way. 7. Add the rinsed tissue to ice-cold CP2 medium making sure to use 40 mL CP2 for every 4–5 g tissue (see Note 4). Stir gently for 5 min on ice to allow protein digestion. 8. Divide the minced tissue into 8–10 Falcon tubes (50-mL) and homogenise it further with a Polytron for 3 s at low speed (see Note 5). Repeat three times for each tube. 9. Return the homogenised tissue to CP2 medium kept on ice and stir for another 6 min. 10. Divide the tissue between 8 plastic 50-mL centrifuge tubes and spin at 490 × g for 10 min (4°C). 11. Filter the supernatant through two layers of muslin, transfer to eight clean 50-mL centrifuge tubes and spin at 10,368 × g for 10 min (4°C). 12. Resuspend the pellets in ice-cold CP1 medium (a few mL for each pellet) and divide into two 50-mL tubes. Make up the volume with CP1 and spin at 10,368 × g for 10 min (4°C). 13. Resuspend the pellets again in CP1, fill the tubes, and spin at 490 × g for 5 min (4°C). 14. Transfer the supernatant to two clean 50-mL centrifuge tubes. Spin at 3,841 × g for 10 min (4°C). 15. Resuspend and combine the final pellets in approximately 1 mL CP1 medium; keep the mitochondrial suspension on ice at all times. 16. Determine the mitochondrial protein concentration (we generally use a biuret assay). This isolation procedure yields typically 40–50 mg mitochondrial protein. 3.2. Preparation of TPMP-Selective Electrode Sleeves

1. Dissolve 1 g polyvinylchloride in 20 mL THF: stir in a covered 50-mL conical flask until fully dissolved, which takes approximately 1 h. 2. Combine 6 mL (10 mM) tetraphenylboron and 20 mL polyvinylchloride (both in THF) in a 100-mL flask; mix vigorously. 3. Add 3 mL dioctylphthalate and continue mixing. 4. Divide this mixture equally into five Petri dishes (10-cm diameter) and allow to dry on a absolutely level surface in a switched-off fume hood (see Note 6). 5. Complete evaporation of the solvent, which will take 24–48 h, yields a colourless and fairly robust membrane that needs to be attached to an electrode sleeve that should be prepared from a

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piece of PVC tubing with a 4-mm outside diameter and approximately 4-cm length; we use domestic earth sleeve from an electrical supplier for this purpose (see Note 7). 6. Put a drop of THF on the membrane and place the PVC sleeve squarely on the drop (alternatively, dip one end of the PVC sleeve in THF and then place on the membrane); support the sleeve until the THF has evaporated and the sleeve is stuck firmly to the membrane. Multiple sleeves can be stuck in each Petri dish; they should be left for 24–48 h to cure. 7. Cut around the sleeve with a very sharp razor blade and remove it, together with an attached membrane patch, from the Petri dish; trim away any excess membrane with scissors. 8. Fill the electrode sleeves with TPMP (10 mM) and check for any leaks; these sleeves should be immersed in TPMP (10 mM) for at least 48 h before use and may be stored dry or in this way for many years. 3.3. Mitochondrial Oxygen Consumption

1. Oxygen electrode assembly – Set up the oxygen electrode following the manufacturer’s instructions (see Note 8). To record dissolved oxygen tensions continuously, connect the electrode controller unit (which applies a polarising voltage across the oxygen electrode’s platinum cathode and Ag/AgCl anode and converts oxygen-induced currents into voltages) to the PowerLab analog-to-digital signal converter that, in turn, should be linked to a personal computer running LabChart software. Less conveniently, a conventional strip chart recorder can be used to record the data instead. 2. Oxygen electrode calibration – When the set-up’s platinum electrode is polarised at −0.6 V with respect to the silver electrode, the oxygen-induced current is linearly dependent on the dissolved oxygen tension and a two-point calibration is sufficient to convert the electrical signal into a biologically meaningful unit. To calibrate, add 3.5 mL air-saturated and pre-heated KHEP to a temperature-controlled (37°C) and well-stirred oxygen electrode chamber, apply the plunger, and record a voltage that reflects the maximum dissolved oxygen tension; this tension has a value of 406 nmol atomic oxygen per mL at 37°C (8). To obtain a zero-oxygen signal, remove the plunger and add a few sodium dithionite crystals, which will remove any dissolved oxygen chemically (see Note 9). Between experimental traces, rinse the oxygen electrode vessel thoroughly with distilled water and ethanol (see Note 10). 3. Calculation of oxygen consumption rates – When isolated mitochondria are incubated with a suitable electron donor, they will exhibit a respiratory activity that is directly proportional to the slope of the time-resolved oxygen trace. LabChart software

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provides a straightforward option to calculate and visualise the temporal derivative of the oxygen progress curves. This software thus facilitates rapid and objective determination of oxygen consumption rates and, importantly, allows accurate judgements as to whether steady respiratory states have been attained. Mitochondrial respiratory rates should be calculated and presented as specific activities normalised to the amount of mitochondrial protein in the assay to allow direct comparison with published activities. 4. Coupling of oxidative phosphorylation – It is generally important to ascertain if and to what extent isolated mitochondria have retained coupling between oxygen consumption and ATP synthesis; mitochondrial samples that have lost this coupling are considered poor physiological models and should not be used for proton leak and some electron leak measurements. Oxygen uptake measurements during the various mitochondrial states defined by Chance and Williams (9) provide insight in the degree of coupling, although a more complete understanding can be obtained from modular kinetic experiments that also take Δψ values into account (cf. Subheading 3.5). Incubate rat skeletal muscle mitochondria (0.35 mg/mL) at 37°C in 3.5 mL KHEP until a stable oxygen electrode signal is achieved (state 1). Then add, sequentially, rotenone (4 μM), succinate (5 mM), and ADP (50–100 μM) to, respectively, inhibit respiratory complex I, and to provide an electron donor and a phosphorylation substrate. Well-coupled mitochondria will exhibit a substantial oxygen consumption rate under these conditions (state 3), which will decrease significantly after a short while when all ADP is depleted (state 4). To provoke further state 3/state 4 transitions more ADP aliquots (50–100 μM) may be added. The experiment comes to an obvious end when all oxygen has been exhausted and an anaerobic state has been reached (state 5). A respiratory control ratio (RCR) can be derived from the recorded trace, which is a parameter defined as the quotient of oxygen uptake rates under state 3 and state 4 conditions. A high coupling efficiency of oxidative phosphorylation will be reflected by a high RCR and we typically observe a value of 4 with rat skeletal muscle mitochondria oxidising succinate. In a similar experiment, state 3 respiratory activity can be inhibited by the ATP synthase inhibitor oligomycin (0.7 μg/mL) and then be re-stimulated by the protonophore FCCP (1.5 μM). The ratio of FCCP-stimulated and oligomycin-inhibited oxygen uptake rates reflects the extent to which mitochondrial electron transfer is coupled to proton translocation. The FCCP-oligomycin rate ratio resembles an RCR as it indicates the intactness of the mitochondrial inner membrane, but it differs in so much that this ratio is not controlled to any extent by the ATP synthase.

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The above experiments may also be performed with a combination of pyruvate and malate to engage respiratory complex I in oxidative phosphorylation. These substrates should both be added at 5 mM and rotenone should be omitted from the assay buffer. 3.4. Mitochondrial Membrane Potential Measurements

1. TPMP electrode assembly – Using an automatic pipette or a microsyringe, add TPMP (10 mM) to a TPMP sleeve filling it to about 1 cm from the top; avoid trapping any air that would prevent proper electrical contact. Attach the filled sleeve to screened cable such that the connected platinum wire sticks into the TPMP solution, but the soldered connection remains dry. A convenient way to achieve a tight fit is to use a yellow pipette tip, with the very end cut off, as a linker; puncture this pipette tip to prevent pressure changes that may damage the membrane during assembly. Insert both the TPMP sleeve and the solid-state reference electrode into the oxygen electrode chamber through an adapted plunger and connect them to a voltmeter (a pH meter is ideal) that is linked to the same PowerLab digital data acquisition system that is used for the oxygen uptake measurements. 2. TPMP electrode conditioning – Add 3.5 mL pre-heated KHEP to a temperature-controlled (37°C) and well-stirred oxygen electrode chamber, insert the TPMP and reference electrodes, and wait until the signal is stable. Using the 1 mM stock solution, add TPMP to a concentration of 1 μM, which should cause a substantial electrode response; increasing the TPMP level with 1 μM increments should result in successively smaller responses. The signal should become stable and virtually noise-free at 5 μM TPMP. It may take several of these conditioning events, and even an overnight incubation in 2–3 μM TPMP, until a sleeve exhibits drift- and noise-free behaviour and it responds to the TPMP dose in a truly logarithmic fashion. Moreover, electrode behaviour tends to improve with use. If such improvement is not observed rapidly, the easiest solution is to try another sleeve; further troubleshooting suggestions are given in (10). Following conditioning and between experiments, TPMP sleeves may be stored in medium containing 2–3 μM TPMP for many months. 3. TPMP electrode calibration and correction for small baseline drift – Although the TPMP electrode exhibits fairly reproducible behaviour within daily sets of experimental traces, we nevertheless recommend to calibrate its response and assess possible drift for each trace individually. So for a typical experiment, incubate rat skeletal muscle mitochondria (0.35 mg/mL) at 37°C in 3.5 mL KHEP. Add rotenone (4 μM) and nigericin (350 ng/mL) to, respectively, inhibit respiratory complex I and collapse the pH gradient across the mitochondrial inner

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membrane. Insert the TPMP and reference electrodes and wait until a stable signal is observed. Using the 1 mM solution, add TPMP with 0.5 μM increments to a concentration of 2.5 μM; await a stable signal before each successive addition. Energise the mitochondrial inner membrane by adding succinate (4 mM). If the mitochondria are well coupled, energisation will result in significant TPMP accumulation into the mitochondrial matrix and, therefore, in a substantial decrease of the electrode signal. Effectors specific to a particular experiment can now be added; in general, wait until a steady signal is obtained before making additions (see Note 11). The experiment is completed by adding FCCP (1.5 μM), which will dissipate the mitochondrial proton-motive force (i.e. Δψ when nigericin is present), will thus cause TPMP release from the mitochondria and will bring the external TPMP concentration back to 2.5 μM within a minute or so. Comparison of the final electrode signal with that observed before succinate addition allows determination of a possible electrode drift rate. If deemed acceptably small, then this rate should be taken into account in the calculation of external TPMP levels (cf. Subheading 3.4). If electrode drift is substantial and/or persistent, its cause(s) should be identified and rectified (cf. Subheading 3.2). 4. Calculation of membrane potentials – Lipophilic cations such as TPMP+ equilibrate across the mitochondrial inner membrane according to the Nernst equation: Δψ = 61.5 log([TPMP+]in/ [TPMP+]out) at 37°C. The membrane potential can thus be calculated from the external (i.e. extra-mitochondrial) and matrix TPMP concentrations. To enable this calculation, measure first the signal deflections from the 2.5 μM TPMP baseline observed at the various applied TPMP levels; plot this deflection as a function of log(TPMP concentration). This should yield a linear calibration relation, which can be used to calculate external TPMP concentrations throughout the experiment. Subtracting the external concentration from the 2.5 μM TPMP applied provides the TPMP level taken up by mitochondria, which is proportional to its concentration in the matrix. To calculate a reliable concentration, however, it is necessary to know how much of the hydrophobic TPMP is actually freely present within the matrix and how much has in fact bound to mitochondrial membranes. Excellent methods to assess probe binding to membranes and other mitochondrial and/or cellular components have been described before (10). For rat skeletal muscle mitochondria, TPMP-binding has been determined as a function of mitochondrial matrix volume and the obtained correction factor is 0.35 mg mitochondrial protein per μL matrix volume (11). Multiplication of this correction factor by the TPMP taken up by mitochondria, normalised to the

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mitochondrial protein present in the assay (mg/μL), yields the free matrix TPMP concentration. Δψ values in rat skeletal muscle mitochondria, incubated at 37°C and 0.00035 mg/μL, can, therefore, be calculated from applied and external TPMP concentrations as: æ ([TPMP + ]applied - [TPMP + ]external ) ´ 0.35 ö ÷ 61.5 log ç ç ÷ 0.00035 ´ [TPMP + ]external è ø 3.5. Proton Leak: A Modular Kinetic Description of Oxidative Phosphorylation

Oxidative phosphorylation is a process that can be divided conceptually into events that either generate or dissipate the mitochondrial membrane potential (12). Such a modular view of mitochondrial energy transduction (Fig. 1a) emphasises that steady-state respiratory activities and Δψ levels result from the kinetic interplay between activity of the mitochondrial electron transfer chain (Δψ-producer) and the combined activities of the ADP phosphorylation machinery and the proton leak across the mitochondrial inner membrane (Δψ-consumers). The kinetic dependency of Δψ-establishing and Δψ-dissipating modules on Δψ is modelled in Fig. 1b based on the assumption that the two Δψ-dissipating modules respond exponentially to Δψ and the Δψestablishing module responds hyperbolically (this is not strictly correct, but serves to illustrate the method here). In mitochondria incubated under phosphorylating conditions, a respiratory steady state is reached when the Δψ-establishing rate is equal to the sum of the Δψ-dissipation rates (Fig. 1b, state 3). Steady states achieved upon either inhibition of phosphorylation with oligomycin or dissipation of the mitochondrial proton-motive force with excess FCCP are labelled “state 4” and “state F”, respectively (Fig. 1b). Note that the oligomycin-inhibited state is generally comparable to the steady state reached in the absence of ADP (state 4). Below, we describe methods to determine modular kinetic relations experimentally. 1. Theoretical principle – The kinetic behaviour of a “Δψproducer” can be established by specific modulation of a “Δψconsumer” (Fig. 1c) and, reciprocally, the behaviour of a consumer is revealed upon modulation of a producer (Fig. 1d; (12)). In other words, if mitochondrial respiration is titrated under state 3 conditions with sub-saturating amounts of an electron transfer chain inhibitor, the kinetics with respect to Δψ of the total Δψ-dissipating activity (i.e. leak + phosphorylation) are revealed. If this titration is performed in the presence of oligomycin, the kinetics of proton leak alone are obtained, which may be subtracted from the overall Δψ-dissipating kinetics to reveal the behaviour of the ADP-phosphorylation module. If respiration is titrated with FCCP, either in the presence or the absence of oligomycin, the kinetic dependency of the

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Fig. 1. Modelled modular kinetics during oxidative phosphorylation in isolated mitochondria. (a) Oxidative phosphorylation considered from a top-down perspective as an interaction between processes that establish Δψ (electron transfer chain, ETC) and those that dissipate it (proton leak and phosphorylation). (b) The kinetic dependency of both the ETC and the sum of proton leak and phosphorylation activities (fluxes expressed in arbitrary units, see Note 9) on Δψ was modelled using exponential (proton leak, phosphorylation) and hyperbolic (ETC) equations. The model assumes zero ΔpH so that the protonmotive force equals Δψ. Steady states are achieved when Δψ-establishing flux equals total Δψ-dissipating flux, which is reflected by intersections of the curves describing the behaviour of the respective activities. Modelled steady states include those reached in the presence (state 3) and the absence of ADP (state 4) as well as that achieved in the presence of FCCP (state F). (c) Titration of oligomycin-insensitive respiratory activity with sub-saturating amounts of FCCP increases proton leak activity specifically. New steady states will thus be achieved that reflect the altered proton leak and the unaffected ETC kinetics (states F1–F3). (d) Titration of succinate oxidation with sub-saturating amounts of malonate inhibits ETC activity specifically. In this case, new steady states will be reached that reflect the altered ETC and the unaffected Δψ-dissipating kinetics (states M1–M3). Malonate titrations under phosphorylating conditions thus reveal the kinetic behaviour of the sum of Δψ-dissipating behaviour and titrations in the presence of oligomycin that of proton leak alone.

respiratory chain on Δψ is established. During these respiration titrations, successive steady states will be attained in which both the oxygen uptake rate (flux) and Δψ (intermediate concentration) should be measured simultaneously. Plotting oxygen consumption rates as a function of the concomitant Δψ values allows empirical construction of modular kinetic plots. 2. Proton leak kinetics – Add rat skeletal muscle mitochondria (0.35 mg/mL) to 3.5 mL KHEP in an oxygen electrode vessel incubated at 37°C in the presence of rotenone (4 μM), nigericin (350 ng/mL), and oligomycin (0.7 μg/mL). Apply the plunger,

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insert the TPMP and reference electrodes, and record both oxygen and TPMP signals. Make five sequential TPMP additions of 0.5 μM each. Add succinate (4 mM) to initiate oligomycinresistant respiration and to energise the mitochondria. When the oxygen uptake rate and TPMP signal have stabilised, make at least five successive malonate additions increasing its concentration gradually to approximately 0.5 mM; finish the trace by adding FCCP (1.5 μM). Calculate specific oxygen consumption rates (see Note 12) and plot as a function of the concomitant Δψ values. The leak rate dependency on Δψ may be modelled approximately by an exponential expression. 3. Phosphorylation kinetics – Perform a trace identical to that described for the proton leak kinetics, but omit oligomycin from the incubation mixture. Instead, include sufficient ADP or an ADP-regenerating system to enable phosphorylating respiration throughout the trace. The data obtained in this experiment represent the kinetic dependency of the sum of phosphorylation and leak activities on Δψ. Subtraction of proton leak kinetics yields the Δψ-dependency of the phosphorylation module alone (see Note 12), which may also be approximated by an exponential expression. 4. Electron transfer chain kinetics – Add rat skeletal muscle mitochondria (0.35 mg/mL) to 3.5 mL KHEP in an oxygen electrode vessel incubated at 37°C in the presence of rotenone (4 μM), nigericin (350 ng/mL), and oligomycin (0.7 μg/mL). Add TPMP to 2.5 μM in five equimolar increments and add succinate (4 mM) to initiate respiration. When the oxygen uptake rate and TPMP signal have stabilised, make at least 5 successive FCCP additions, increasing its concentration gradually to approximately 0.8 μM; finish the trace by adding surplus FCCP (1.5 μM). Calculate specific oxygen consumption rates and plot as a function of the concomitant Δψ values. The electron transfer chain rate dependency on Δψ may be modelled approximately by a hyperbolic expression. Simultaneous measurement of oxygen consumption and Δψ provides considerably more insight in oxidative phosphorylation than individual determination of these parameters. (1) Mitochondrial coupling efficiency is better understood from modular kinetic information than from respiration alone since the kinetic behaviour of the respective modules explains the exact nature of a particular RCR (cf. Subheading 3.3, step 4) and, moreover, identifies the underlying cause(s) for possible differences in RCRs between experimental conditions and systems. (2) Mitochondrial proton leak activity can be compared in a meaningful manner between experimental systems and conditions as the oligomycin-insensitive respiratory rates can be corrected for possible differences in the leak’s driving force, i.e. Δψ; similarly, activity of the other kinetic

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modules can be probed at identical Δψ values. (3) Site(s) at which suspected effectors of oxidative phosphorylation act can be identified unambiguously. (4) Conclusive evidence can be obtained as to the role played by poorly characterised proteins in oxidative phosphorylation. (5) Elasticities of all kinetic modules to Δψ can be calculated, which allows control and regulation of oxidative phosphorylation to be quantified (13). 3.6. Electron Leak

1. Principle of the Amplex Red assay – Electron leaks during oxidative phosphorylation result in the single electron reduction of molecular oxygen to superoxide, a reaction that occurs predominantly at respiratory complexes I and III. Assays exist to measure superoxide directly in isolated mitochondria (14) but are of limited use as they only detect superoxide that is released from the mitochondria, which is due to the impermeability of the mitochondrial inner membrane to most charged molecular species. This means that these direct assays only report superoxide that is produced at the cytoplasmic side of the inner membrane and fail to reveal any production that is directed towards or indeed occurs within the mitochondrial matrix. However, matrix superoxide is converted rapidly to H2O2 by endogenous superoxide dismutases and H2O2 is an uncharged molecule that will diffuse freely across mitochondrial membranes. In isolated mitochondria, released H2O2 can be detected readily through its horseradish peroxidasecatalysed reaction with the fluorogenic probe Amplex Red (N-acetyl-3,7-dihydroxyphenoxazine). As superoxide production and H2O2 release rates are proportional, the Amplex Red assay provides a convenient way to determine electron leak activities in isolated mitochondria. It should be noted, however, that the assay is only semi-quantitative unless specifically corrected: not all matrix-formed H2O2 will diffuse to the medium, as part is detoxified by endogenous mitochondrial hydrogen peroxidases, and these losses need to be accounted for (15). 2. Spectrofluorometer set-up and general assay conditions – Set the spectrofluorometer’s excitation wavelength to 563 nm, the emission wavelength to 587 nm and adjust the assay temperature to 37°C. Carefully add 2 mL KHE (pre-heated at 37°C) to a stirred cuvette avoiding air bubbles in the light path. Add mitochondria to a final concentration of 0.35 mg/mL, Amplex Red to 50 μM, horseradish peroxidase to 6 U/mL and superoxide dismutase to 30 U/mL (see Note 13). Start recording data in a time-resolved manner to establish possible background fluorescence and initiate the experiment by adding an appropriate respiratory substrate; inhibitors and other effectors may now be added as required (see below for specifics). This will yield a fluorescence progress curve similar to the one shown in Fig. 2a.

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3. Calibration – To convert fluorescence units into moles of H2O2, a separate calibration curve is generated for each applied experimental condition. This is necessary to account for possible quenching or background effects of particular assay components. Set up an assay as described above and include all components except for the respiratory substrate. Add 5 equimolar amounts of H2O2 (1 μL additions of a 1 mM stock) in a stepwise fashion (Fig. 2b) and plot the observed fluorescence as a function of the added H2O2 amount (Fig. 2c). The rate of H2O2 production (nmol/min/mg) under a particular experimental condition is calculated by dividing the slope of the fluorescence trace (fluorescence units/min) by the slope of the corresponding standard curve (fluorescence units/nmol H2O2) and the total amount of protein (mg) in

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the cuvette. When using new reagents or conditions, it is essential to perform controls without mitochondria to assess the magnitude of any non-mitochondrial interferences. 4. Superoxide production by complex I – Respiratory complex I (NADH:ubiquinone oxidoreductase) can produce superoxide when it is reduced by the oxidation of either NADH or ubiquinol. In other words, this superoxide production may be observed when isolated mitochondria are incubated under conditions that allow forward or reverse electron transfer, respectively. To measure “forward” superoxide production, set up an assay as described above and add a respiratory substrate that leads to a substantial reduction of the mitochondrial NAD+/NADH pool, for example a combination of malate and glutamate both added at 5 mM. Superoxide is believed to be produced under these conditions at the flavin site of complex I (16), and its production rate is increased substantially by including 4 μM rotenone in the assay to block efflux of electrons from the complex. It should be noted, however, that it is not clear whether or not all this “forward” superoxide arises at complex I, since a high reduction of the NAD+/NADH pool may also favour superoxide production by α-ketoglutarate dehydrogenase (17). To measure “reverse” superoxide production, set up an assay as described above and add 5 mM succinate to initiate the experiment. The so-established reduced ubiquinone pool and high proton-motive force favour reduction of complex I via ubiquinol oxidation and result in substantial rates of rotenone-sensitive superoxide production that can be modulated by the pH gradient across the mitochondrial inner membrane. This “reverse” superoxide is thought to arise from another site than the flavin, perhaps the ubiquinone-binding site of complex I (18). 5. Superoxide production by complex III – Complex III (the cytochrome bc1 complex) oxidises ubiquinone molecules that are reduced by substrate oxidation and it produces superoxide when the proton-motive force is high or in the presence antimycin A, an inhibitor of ubiquinone reduction by complex III in the Q-cycle mechanism. Any respiratory substrate that reduces the quinone pool may be used to feed electrons into complex III. To measure superoxide production from complex III, set up an assay as described above and add either succinate (5 mM) or a combination of malate/glutamate (both at 5 mM) to initiate the experiment and subsequently add antimycin A (2 μM) to achieve maximum rates. Superoxide production that is inhibited by stigmatellin under these conditions emanates from complex III (see Note 13).

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4. Notes 1. The oxygen-permeable Teflon membrane supplied by Rank Brothers may be substituted with high-sensitivity membrane (YSI Life Sciences, Ohio, USA) to decrease the response time of the oxygen electrode. 2. Bovine serum albumin should be added to the KHEP assay buffer on the day of use; sprinkle the required amount (300 mg/100 mL) on top of the solution and let it dissolve without stirring to prevent clotting. 3. H2O2 working stocks (1 mM) must be prepared fresh on the day of use because these dilute solutions are unstable. 4. The ratio of tissue to CP2 medium is critically important because it sets the appropriate conditions for protease-mediated muscle digestion. Too much digestion will result in poor mitochondrial quality, whereas too little digestion will result in a low yield. 5. The Polytron’s speed and sharpness affects mitochondrial yield and quality: too fast homogenisation will result in damaged mitochondria whilst a slower setting will lower the yield. The ideal speed setting varies for different Polytron types and needs to be determined empirically. 6. Modern fume hoods may sometimes not be switched off; to prevent hasty evaporation and surface rippling of the membrane, cover the Petri dishes in this case with lids but leave small openings to enable drying. 7. When cutting PVC tubing, it is pivotal to get a very clean and straight cut as the sleeve will not stick to the membrane otherwise. The likelihood of obtaining such cuts is maximised when performed by two persons: one pulls the tubing as tightly as possible and another cuts it very quickly and cleanly with a sharp razor blade. 8. Whether or not the assembled set-up detects oxygen and responds rapidly to changes in oxygen concentration can be checked conveniently by switching off the electronic stirrer temporarily, which should cause a substantial and fast signal deflection. Lack of stirring means that oxygen levels near the platinum cathode and the bulk assay medium are no longer kept in rapid equilibrium and, therefore, that the oxygen uptake at the cathode will lead quickly to localised oxygen deprivation. 9. Sodium dithionite cannot always be used to deplete oxygen from the assay buffer as it may interfere with other measurements such as the detection of TPMP. An alternative way to obtain the zero-oxygen signal is to rely on mitochondrial

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respiration to consume all oxygen and use the anaerobic signal achieved at the end of an experimental trace. If this “biological zero” is known from previous experiments to be similar to the signal obtained from an unpolarised electrode, this “electrical zero” could be used as a quick-and-dirty approximation of a zero-oxygen signal. 10. Some respiratory effectors (e.g. rotenone and FCCP) are hard to wash away between experimental traces as they tend to stick to the plastic oxygen electrode vessel. Even when a glass vessel is used to reduce this sticking issue, it may sometimes be wise to rinse the electrode chamber between traces with a concentrated (3% w/v) bovine serum albumin solution or with a suspension of frozen-thawed mitochondria that were left over from earlier experiments. Bovine serum albumin and mitochondria will bind all possible nasties and using them as a mop will greatly reduce the likelihood of effector carry-over. 11. Once in a while, the TPMP electrode will exhibit small drift. It is important, however, to distinguish such drift from a lack of biological steady state as it is possible that Δψ is changing slowly because of alteration in electron transfer chain, proton leak and/or phosphorylation activities. Changes in these processes, however, will also likely be reflected in altered oxygen consumption rates. An important indication, therefore, that a slowly changing TPMP signal reflects biological reality and is not merely due to electrode drift, is a concomitantly changing oxygen electrode signal. 12. Oligomycin-insensitive oxygen uptake activities are converted to proton flux rates by multiplying them with appropriate H+/O ratios, which are 6 and 10 during the oxidation of succinate and pyruvate/malate, respectively. ADP-phosphorylation rates are obtained by multiplying the leak-corrected state 3 respiratory activities with the appropriate P/O ratios, which are 1.39 and 2.31 during the oxidation of succinate and pyruvate/ malate, respectively (2). 13. Superoxide produced by complex III is released to both the cytoplasmic and the matrix side of the mitochondrial inner membrane. Superoxide dismutase should, therefore, always be present in the assay medium to ensure accurate estimation of the total superoxide produced.

Acknowledgments We thank Julie Buckingham for useful advice on the preparation of TPMP sleeves.

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References 1. Nicholls DG, Ferguson SJ (2002) Bioenergetics 3. Academic Press, London 2. Brand MD (2005) The efficiency and plasticity of mitochondrial energy transduction. Biochem Soc Trans 33:897–904 3. Brand MD, Affourtit C, Esteves TC, Green K, Lambert AJ, Miwa S, Pakay JL, Parker N (2004) Mitochondrial superoxide: production, biological effects, and activation of uncoupling proteins. Free Radic Biol Med 37: 755–767 4. Affourtit C, Brand MD (2008) Uncoupling protein-2 contributes significantly to high mitochondrial proton leak in INS-1E insulinoma cells and attenuates glucose-stimulated insulin secretion. Biochem J 409:199–204 5. Dröge W (2002) Free radicals in the physiological control of cell function. Physiol Rev 82:47–95 6. Ashour B, Hansford RG (1983) Effect of fatty acids and ketones on the activity of pyruvate dehydrogenase in skeletal-muscle mitochondria. Biochem J 214:725–736 7. Letellier T, Malgat M, Mazat JP (1993) Control of oxidative phosphorylation in rat muscle mitochondria: implications for mitochondrial myopathies. Biochim Biophys Acta 1141: 58–64 8. Reynafarje B, Costa LE, Lehninger AL (1985) O2 solubility in aqueous media determined by a kinetic method. Anal Biochem 145: 406–418 9. Chance B, Williams GR (1955) Respiratory enzymes in oxidative phosphorylation. III. The steady state. J Biol Chem 217:409–427 10. Brand MD (1995) Measurement of mitochondrial protonmotive force. In: Brown GC,

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Cooper CE (eds) Bioenergetics: a practical approach. IRL, Oxford, pp 39–62 Rolfe DFS, Hulbert AJ, Brand MD (1994) Characteristics of mitochondrial proton leak and control of oxidative phosphorylation in the major oxygen-consuming tissues of the rat. Biochim Biophys Acta 1188:405–416 Brand MD (1998) Top-down elasticity analysis and its application to energy metabolism in isolated mitochondria and intact cells. Mol Cell Biochem 184:13–20 Brand MD (1996) Top down metabolic control analysis. J Theor Biol 182:351–360 Turrens JF, Boveris A (1980) Generation of superoxide anion by the NADH dehydrogenase of bovine heart mitochondria. Biochem J 191:421–427 Treberg JR, Quinlan CL, Brand MD (2010) Hydrogen peroxide efflux from muscle mitochondria underestimates matrix superoxide production: a correction using glutathione depletion. FEBS J 277(13):2766–2778 Kussmaul L, Hirst J (2006) The mechanism of superoxide production by NADH:ubiquinone oxidoreductase (complex I) from bovine heart mitochondria. Proc Natl Acad Sci USA 103: 7607–7612 Starkov AA, Fiskum G, Chinopoulos C, Lorenzo BJ, Browne SE, Patel MS, Beal MF (2004) Mitochondrial alpha-ketoglutarate dehydrogenase complex generates reactive oxygen species. J Neurosci 24:7779–7788 Lambert AJ, Brand MD (2004) Inhibitors of the quinone-binding site allow rapid superoxide production from mitochondrial NADH: ubiquinone oxidoreductase (complex I). J Biol Chem 279:39414–39420

Chapter 12 Relation Between Mitochondrial Membrane Potential and ROS Formation Jan M. Suski, Magdalena Lebiedzinska, Massimo Bonora, Paolo Pinton, Jerzy Duszynski, and Mariusz R. Wieckowski Abstract Mitochondria are considered as the main source of reactive oxygen species (ROS) in the cell. For this reason, they have been recognized as a source of various pathological conditions as well as aging. Chronic increase in the rate of ROS production is responsible for the accumulation of ROS-associated damages in DNA, proteins, and lipids, and may result in progressive cell dysfunctions and, in a consequence, apoptosis, increasing the overall probability of an organism’s pathological conditions. The superoxide anion is the main undesired by-product of mitochondrial oxidative phosphorylation. Its production is triggered by a leak of electrons from the mitochondrial respiratory chain and the reaction of these electrons with O2. Superoxide dismutase (MnSOD, SOD2) from the mitochondrial matrix as well as superoxide dismutase (Cu/ZnSOD, SOD1) present in small amounts in the mitochondrial intramembrane space, convert superoxide anion to hydrogen peroxide, which can be then converted by catalase to harmless H2O. In this chapter, we describe a relation between mitochondrial membrane potential and the rate of ROS formation. We present different methods applicable for isolated mitochondria or intact cells. We also present experiments demonstrating that a magnitude and a direction (increase or decrease) of a change in mitochondrial ROS production depends on the metabolic state of this organelle. Key words: Mitochondria, Membrane potential, ROS, Amplex Red, JC-1, Superoxide, Respiration, Confocal microscopy

1. Introduction It has been repeatedly demonstrated, on different experimental models, that a strong positive correlation exists between mitochondrial membrane potential (DY) and reactive oxygen species (ROS) production (1, 2). At present, it is widely accepted that

Jan M. Suski and Magdalena Lebiedzinska contributed equally to prepare this chapter. Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_12, © Springer Science+Business Media, LLC 2012

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mitochondria produce more ROS at high membrane potential. It has been shown that ROS production dramatically increases above 140 mV (2). Studies performed on mitochondria from Drosophila melanogaster showed that even a slight decrease in the DY (10 mV) can cause a significant decrease in ROS production (according to authors the decrease of ROS production diminished by approximately 70%) by complex I of the respiratory chain (3). In contrast, an increase in the DY produced either by a closure of the mitochondrial permeability transition pore or an inhibition of ATP synthase (4) is associated with increased ROS production. Interestingly, in certain pathological conditions, opposite correlations between DY and ROS production can also be observed. In the case of ATP synthase dysfunction (mutation T8993G in the mitochondrial ATPase-6 gene), higher DY and increased ROS production is observed (5). On the other hand, in the case of mitochondrial disorders associated with the dysfunctions of the respiratory chain components, lower DY and decreased activity of the respiratory chain is observed with a simultaneous increase in ROS production (6).

2. Materials 2.1. Isolation of Crude Mitochondria from Mouse Brain

1. Stirrer motor with electronic speed controller. 2. Motor-driven tightly fitting glass/Teflon Potter Elvehjem homogenizer. 3. Loose fitting glass/Teflon Potter Elvehjem homogenizer. 4. Homogenization buffer (320 mM sucrose, 10 mM Tris–HCl, 1 mM EDTA, pH 7.4). Store at 4°C (see Note 1). 5. Mitochondria isolation buffer (75 mM sucrose, 225 mM mannitol, 5 mM Tris–HCl, pH 7.4). Store at 4°C (see Note 2).

2.2. Cell Culture

1. Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (heat inactivated), 2 mM L -glutamine, and 1.2% antibiotic: penicillin/streptomycin. 2. KRB saline (135 mM NaCl, 5 mM KCl, 0.4 mM KH2PO4, 1 mM MgSO4, 20 mM HEPES, bring to pH 7.4 with NaOH). Glucose 1 g/L and 1 mM CaCl2 should be added the day of experiment. 3. Fibroblasts: (a) Primary culture of human skin fibroblasts grown from explants of skin biopsies of control individual and patient with mitochondrial disorder. (b) NHDF, neonatal dermal fibroblasts (Lonza).

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4. Ehrlich ascites tumor cells were cultivated in Swiss albino mice and harvested as described in ref. (7). 5. 24-Well Cell Culture Cluster. 2.3. Measurement of Hydrogen Peroxide Production in Isolated Brain Mitochondria

1. Spectrofluorimeter. 2. Measurement medium (75 mM sucrose, 225 mM mannitol, 5 mM Tris–HCl, pH 7.4). Store at 4°C (see Note 2). 3. 0.5 M Succinate, pH 7.4 (adjusted with KOH). 4. 1 mM Antimycin A (ethanol solution). 5. 1 mM Carbonyl cyanide m-chloro phenyl hydrazone (CCCP) (ethanol solution). 6. 5 mM 5-(and-6)-chloromethyl-2¢,7¢-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA), in DMSO.

2.4. Measurement of Superoxide Production by Peroxidase/Amplex Red Assay in Isolated Mitochondria

1. Multiwell plate reader. 2. 24-Well Cell Culture Cluster. 3. Reaction buffer (75 mM sucrose, 225 mM mannitol, 5 mM Tris–HCl, pH 7.4). Store at 4°C (see Note 2). 4. 5 mM Amplex Red. 5. 7 U/ml Peroxidase. 6. 0.5 M Glutamate, pH 7.4. 7. 0.5 M Malate, pH 7.4. 8. 1 mM Oligomycin (ethanol solution). 9. 1 mM CCCP (ethanol solution). 10. 1 mM Antimycin A (ethanol solution).

2.5. Measurement of the Mitochondrial Transmembrane Potential Using Safranine O in Isolated Mouse Brain Mitochondria

1. Spectrofluorimeter. 2. Measurement medium (75 mM sucrose, 225 mM mannitol, 5 mM Tris–HCl, pH 7.4). Store at 4°C (see Note 2). 3. 5 mM Safranine O. 4. 0.5 M Succinate, pH 7.4 (adjusted with KOH). 5. 1 mM Antimycin A (ethanol solution). 6. 1 mM Oligomycin (ethanol solution). 7. 1 mM CCCP (ethanol solution).

2.6. Measurement of the Mitochondrial Transmembrane Potential in Human Fibroblasts Using JC-1

1. Multiwell plate reader. 2. 24-Well Cell Culture Cluster. 3. 5 mM JC1 (5,5¢,6,6¢-tetrachloro-1,1¢,3,3¢-tetraethylbenzimidazolylcarbocyanine iodide); (Invitrogen/Molecular Probes) in DMSO. Store at −20°C. 4. KRB saline (see item 2 in Subheading 2.2).

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5. 1 mM Antimycin A (ethanol solution). 6. 1 mM Oligomycin (ethanol solution). 2.7. Measurement of “Mitochondrial Matrix” Superoxide Production in Human Fibroblasts Using MitoSOX Red

1. Multiwell plate reader. 2. 24-Well Cell Culture Cluster. 3. 5 mM MitoSox Red in DMSO. Store at −20°C. 4. KRB saline (see item 2 in Subheading 2.2). 5. 1 mM Antimycin A (ethanol solution). 6. 1 mM Oligomycin (ethanol solution).

2.8. Measurement of the “Cytosolic” Superoxide Production in Human Fibroblasts Using DHE

1. Multiwell plate reader. 2. 24-Well Cell Culture Cluster. 3. 10 mM Dihydroethidium (hydroethidine) in DMSO. Store at −20°C. 4. KRB saline (see item 2 in Subheading 2.2). 5. 1 mM Antimycin A (ethanol solution). 6. 1 mM Oligomycin (ethanol solution).

2.9. Measurement of the Oxygen Consumption in Ehrlich Ascites Tumor Cells

1. Clark-type oxygen electrode (YSI, Yellow Springs, OH, USA), equipped with a unit calculating the equivalent to the rate of oxygen consumption (first derivative of the oxygen concentration trace). 2. KRB saline (see item 2 in Subheading 2.2). 3. 1 mM Oligomycin (ethanol solution). 4. 1 mM Cyclosporin A (ethanol solution). 5. 1 mM carbonylcyanide p-trifluoromethoxyphenylhydrazone (FCCP) (ethanol solution).

2.10. Fluorometric Measurement of the Mitochondrial Membrane Potential Using TMRM in Ehrlich Ascites Tumor Cells

1. Spectrofluorometer. 2. 100 mM TMRM in H2O. 3. KRB saline (see item 2 in Subheading 2.2). 4. 1 mM Oligomycin (ethanol solution). 5. 1 mM Cyclosporin A (ethanol solution). 6. 1 mM FCCP (ethanol solution).

2.11. Fluorometric Measurement of Hydrogen Peroxide Production in Ehrlich Ascites Tumor Cells

1. Spectrofluorometer. 2. 5 mM CM-H2DCFDA in DMSO. 3. KRB saline (see item 2 in Subheading 2.2). 4. 1 mM Oligomycin (ethanol solution). 5. 1 mM Cyclosporin A (ethanol solution). 6. 1 mM FCCP (ethanol solution).

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1. Laser scanning or Spinning disk confocal microscope equipped with 546 or 561 nm laser. 2. Microscope cover slips (24 mm diameter, 0.15 mm thickness). 3. KRB saline (see item 2 in Subheading 2.2). 4. 10 mM TMRM in absolute ethanol.

2.13. Measurement of Hydrogen Peroxide Production in HeLa Cells Using a Confocal Microscope

1. Laser scanning or Spinning disk confocal microscope equipped with 488nm laser. 2. Microscope cover slips (24 mm diameter, 0.15 mm thickness). 3. KRB saline (see item 2 in Subheading 2.2). 4. 5 mM CM-H2DCFDA in DMSO.

2.14. Measurement of Mitochondrial Superoxide Production in HeLa Cells Using a Confocal Microscope

1. Laser scanning or Spinning disk confocal microscope equipped with 514nm laser.

2.15. Measurement of Cytosolic Calcium in HeLa Cells Using a Confocal Microscope

1. Laser scanning or Spinning disk confocal microscope equipped with 488nm laser.

2. Microscope cover slips (24 mm diameter, 0.15 mm thickness). 3. KRB saline (see item 2 in Subheading 2.2). 4. 5 mM MitoSOX Red in DMSO.

2. Microscope cover slips (24 mm diameter, 0.15 mm thickness). 3. KRB saline (see item 2 in Subheading 2.2). 4. 5 mM Cell-permeant fluo-3 AM in DMSO.

2.16. Measurement of the Respiratory Chain Activity in Human Fibroblasts

1. Multiwell plate reader. 2. 24-Well Cell Culture Cluster. 3. 1 mM Resazurin (7-Hydroxy-3H-phenoxazin-3-one-10-oxide sodium salt) in H2O. Store at −20°C. 4. KRB saline (see item 2 in Subheading 2.2). 5. 1 M KCN in H2O.

2.17. Fluorometric Measurement of Hydrogen Peroxide Production in Human Fibroblasts

1. Multiwell plate reader.

2.18. Determination of Protein Concentration

1. Lysis buffer (50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 1% Triton-X, 0.1% SDS, and 1% sodium deoxycholate).

2. 24-Well Cell Culture Cluster. 3. 5 mM CM-H2DCFDA in DMSO. Store at −20°C. 4. KRB saline (see item 2 in Subheading 2.2).

2. Bio-Rad Protein Assay. 3. Spectrophotometer. 4. Acryl Cuvettes 10 × 10 × 48 mm.

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3. Methods To study the relationship between mitochondrial ROS production and mitochondrial bioenergetic parameters (like membrane potential and respiration), a variety of methods can be used. Some of them, particularly those dedicated to human fibroblasts could potentially be adapted as diagnostic procedures in case of suspected mitochondrial disorders (5). 3.1. Isolation of Crude Mouse Brain Mitochondria for Measurement of DY and ROS Production

1. Kill the mouse by decapitation, remove the brain immediately, and cool it down at 4°C in the homogenization medium (see Note 3). 2. Wash the brain with the homogenization medium (to remove blood). Add fresh homogenization medium in a proportion of 5 ml/g of brain. 3. Homogenize the brain in a glass Potter–Elvehjem homogenizer with a motor-driven Teflon pestle (see Note 4). 4. Centrifuge the homogenate for 3 min at 1,330 × g at 4°C. 5. Discard the pellet and centrifuge the supernatant for 10 min at 21,000 × g at 4°C. 6. Gently resuspend the resulting mitochondrial pellet in approximately 15 ml of the mitochondria isolation buffer and centrifuge it again for 10 min at 21,000 × g at 4°C. 7. Gently resuspend the final crude mitochondrial pellet in 1–2 ml (depending on the pellet volume) of the mitochondria isolation buffer using a loose Potter–Elvehjem homogenizer. 8. The material can now be used for further experiments (Such isolated mitochondria contain synaptosomes. If necessary additional steps in the isolation procedure can be undertaken to isolate the pure mitochondrial fraction).

3.2. Measurement of Hydrogen Peroxide Production with the Use of CM-H2DCFDA in Isolated Mitochondria

1. Adjust the fluorometer: Excitation – 513 nm; Emission – 530 nm; Slits (excitation and emission) ~ 3. 2. Fill the fluorometer cuvette with 3 ml of the measurement medium (75 mM sucrose, 225 mM mannitol, 5 mM Tris–HCl, pH 7.4. Store at 4°C (see Note 2)) containing 1 mg of mitochondrial protein. 3. Start measurement. 4. Add CM-H2DCFDA to the final 5 mM concentration and record changes in fluorescence. 5. Make traces in the presence of following additions: (a) First trace: 2 mM antimycin A. (b) Second trace: 2 mM FCCP. 6. An example of the results obtained is shown in Fig. 1a.

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Fig. 1. Effect of antimycin A, CCCP, and oligomycin on mitochondrial membrane potential and ROS formation measured in isolated mitochondria. (a) Effect of antimycin A and CCCP on mitochondrial H2O2 production measured with the use of CM-H2DCFDA (Subheading 3.2); (b) Effect of oligomycin, CCCP, and antimycin A on superoxide production measured by peroxidase/Amplex Red assay (Subheading 3.3); (c) Effect of oligomycin, antimycin A, and CCCP on DY measured with the use of Safranine O (Subheading 3.4). Addition of antimycin A or CCCP leads to the collapse of the DY (panel c); however, each compound results in contradictory effects in terms of ROS formation in isolated mitochondria. These effects can be observed with the use of different probes, such as CM-H2DCFDA (panel a) and peroxidase/Amplex Red assay (panel b). Antimycin A is an inhibitor of complex III of the respiratory chain and causes the accumulation of reduced intermediates leading to the chain’s blockage and an increased leakage of electrons. This in turn results in an increase in ROS formation. CCCP as a mitochondrial uncoupler decreases the amount of reduced respiratory chain intermediates and decreases ROS formation (panels a and b). In “coupled” mitochondria, in the absence of ADP, no significant effect of oligomycin (an inhibitor of the mitochondrial ATP synthase) neither on the DY nor on the ROS production is observed (panels b and c). A significant effect of oligomycin on both parameters can be seen in the intact cell model (see Fig. 2).

3.3. Mitochondrial Superoxide Measurement by Peroxidase/Amplex Red Assay

1. Prepare 13 ml of the reaction buffer (see Note 5). 2. Supplement the reaction buffer with 3 ml of Amplex Red, 13 ml of proxidase, 130 ml of glutamate, and 130 ml of malate. 3. Aliquot 0.5 ml of the reaction solution into the wells. 4. Supplement individual wells with oligomycin, FCCP, and antimycin A. 5. Start the reaction with the addition of 100 mg of mitochondrial protein. 6. Place the plate in the microplate reader and read the fluorescence at 510 nm excitation and 595 nm emission wavelengths. An example of the results obtained is shown in Fig. 1b.

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3.4. Measurement of the Mitochondrial Membrane Potential with the Use of Safranine O

1. Adjust the fluorometer: Excitation – 495 nm; Emission – 586 nm; Slits (excitation and emission) ~ 3. 2. Fill the fluorometer cuvette with 3 ml of the measurement medium (see Note 5). 3. Add 3 ml of safranine O and 30 ml of succinate. 4. Start measuring fluorescence. 5. Add the mitochondrial suspension corresponding to about 1 mg of protein and observe changes in fluorescence. 6. Make traces with the following consecutive additions: (a) First trace: 5 ml oligomycin, 5 ml CCCP. (b) Second trace: 5 ml antimycin A. (c) Third trace: 5 ml CCCP. 7. An example of the results obtained is shown in Fig. 1c.

3.5. Measurement of Mitochondrial Membrane Potential in Human Fibroblasts with the Use of JC-1

1. Remove the culture medium and wash the cells gently with warm KRB before preincubation with effectors. Dilute stock solutions of antimycin A and oligomycin to final concentration 2 mM in KRB. Add 0.5 ml of KRB with particular effector to selected wells, and KRB alone to control wells. Preincubate the plate, in the incubator, for 15–30 min prior to the measurement (see Note 6). 2. Prepare 5 mM solution of JC-1 in KRB (see Note 7). 3. Remove the preincubation solution from wells and add the KRB solution containing JC-1. As the effectors must be present till the end of measurement, add adequate amount of each to selected wells (see Notes 5 and 8). 4. Incubate the plate for 10 min in the incubator (see Note 9). 5. Gently wash the cells twice with KRB. 6. Add 0.5 ml of KRB to each well (see Note 10). 7. Place the plate in reader and read the fluorescence. The read must be done twice, first at 485 nm excitation and 520 nm emission wavelengths for JC-1 green fluorescence detection and second at 535 nm excitation and 635 nm emission wavelengths for JC-1 red fluorescence detection. The value of mitochondrial potential is the ratio of red to green fluorescence (see Note 11). An example of the results obtained is shown in Figs. 2a, b and 8a.

3.6. Measurement of Mitochondrial Superoxide Production in Human Fibroblasts with the use of MitoSox Red

1. Remove the culture medium and wash the cells gently with warm KRB before preincubation with effectors. Dilute stock solutions of antimycin A and oligomycin to a final concentration of 2 mM in KRB. Add 0.5 ml of KRB with particular effector to selected wells and KRB alone to control wells. Preincubate the plate, in the incubator, for 15–30 min prior to the measurement (see Note 6).

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2. Prepare 5 mM solution of MitoSox in KRB. Protect the solution from light (see Notes 5 and 8). 3. Remove the preincubation solution from wells and add 0.5 ml of the MitoSox Red solution per well. Add proper amounts of antimycin A or oligomycin to the selected wells (see Notes 5, 8, and 10). 4. Incubate for 10 min in the incubator. 5. Gently wash the cells twice with warm KRB. 6. Add 0.5 ml KRB to each well (see Note 10). 7. Place the plate in the microplate reader and read the fluorescence at 510 nm excitation and 595 nm emission wavelengths (see Note 11). An example of the results obtained is shown in Figs. 2a, b and 8d.

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3.7. Measurement of Cytosolic Superoxide Production in Human Fibroblasts with the Use of DHE

1. Remove the culture medium and wash the cells gently with warm KRB before preincubation with effectors. Dilute stock solutions of antimycin A and oligomycin to a final concentration of 2 mM in KRB. Add 0.5 ml of KRB with particular effector to selected wells and KRB alone to control wells. Preincubate the plate, in the incubator, for 15–30 min prior to the measurement (see Note 6). 2. Prepare 5 mM DHE solution in KRB (see Notes 5, 8, and 10). 3. Remove the preincubation solution from wells and add 0.5 ml of DHE solution to each well of 24-wells plate. As the effectors must be present till the end of measurement, add proper amounts of antimycin A or oligomycin to the selected wells. 4. Incubate for 20 min in the incubator. 5. Gently wash the cells twice with KRB. 6. Add KRB to each well (see Note 10). 7. Place the plate in the microplate reader and read the fluorescence at 535 nm excitation and 635 nm emission wavelengths. This is a single read, no kinetic should be defined. An example of the results obtained is shown in Figs. 2a, b and 8c.

3.8. Measurement of Oxygen Consumption (Respiration) in Ehrlich Ascites Tumor Cells

1. Add cells (approximately 6 mg) to the chamber, fill the chamber with the measurement medium (NaCl 135 mM, KCl 5 mM, KH2PO4 0.4 mM, MgSO4 1 mM, HEPES 20 mM, adjusted to pH 7.4 with NaOH. Glucose 1 g/L and 1 mM CaCl2) and close the chamber. 2. Start the oxygen consumption measurement (first derivative of the oxygen concentration trace). 3. Wait for the stable signal (usually it takes about 1–3 min). 4. Make traces with the following additions: (a) First trace: 1 ml of oligomycin → 1 ml of CsA → 1 ml of FCCP. (b) Second trace: 1 ml of CsA → 1 ml of oligomycin → 1 ml of FCCP. 5. An example of the results obtained is shown in Fig. 3a, b.

3.9. Fluorometric Measurement of Mitochondrial Membrane Potential in Ehrlich Ascites Tumor Cells with the Use of TMRM

1. Adjust the fluorometer: Excitation – 556 nm; Emission – 576 nm; Slits (excitation and emission) ~ 3. 2. Fill the fluorometer cuvette with 3 ml of the measurement medium (NaCl 135 mM, KCl 5 mM, KH2PO4 0.4 mM, MgSO4 1 mM, HEPES 20 mM, adjusted to pH 7.4 with NaOH. Glucose 1 g/L and 1 mM CaCl2) containing 1 × 107 of EAT cells. 3. Add 5 ml of 200 mM TMRM. 4. Start the fluorescence measurement.

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Fig. 3. Effect of oligomycin, cyclosporine A, and FCCP on oxygen consumption, mitochondrial membrane potential, and H2O2 production measured in Ehrlich ascites tumor cells. (a, a¢) oxygen consumption was measured with the use of a Clark-type oxygen electrode (Subheading 3.8) (b, b¢) DY was measured with the use of TMRM in a Shimadzu Spectrofluorimeter RF 5000 (Subheading 3.9); (c) H2O2 production was measured with the use of CM-H2DCFDA in Shimadzu Spectrofluorimeter RF 5000 (Subheading 3.10). The addition of oligomycin (oligo) to the Ehrlich ascites tumor cells caused a decrease in oxygen consumption (manifested as a decrease of first derivative value of the oxygen concentration trace) (Panels a and b) and an increase in mt DY (Panels a¢ and b¢), corresponding to the resting-state (respiratory State 4) level. The effect of oligomycin on mitochondrial bioenergetic parameters is manifested in an increased rate of H2O2 production (Panel c). Further addition of FCCP resulted in the acceleration of oxygen consumption, a rapid collapse of the DY, and decreased rate of H2O2 production. Cyclosporine A (CsA), an inhibitor of the permeability transition pore, added before oligomycin also partially decreased oxygen consumption (Panel b), increased the mt DY (Panel b¢), and increased the rate of H2O2 production (Panel c). Incompatibility of the amplitude changes in TMRM fluorescence with the alterations in the rate of oxygen consumption after oligomycin and cyclosporine A addition was evoked by the effect of both of these compounds on the multidrug resistant (MDR) proteins. Basing on the oxygen consumption data, it is recommended to use an inhibitor of MDR proteins as, e.g.: Sulfinpyrazone 100 mM.

5. Make traces with the following additions: (a) First trace: 5 ml oligomycin, 5 ml of CsA, and 5 ml FCCP. (b) Second trace: 5 ml CsA, 5 ml of oligomycin, and 5 ml FCCP. 6. An example of the results obtained is shown in Fig. 3a¢, b¢. 3.10. Fluorometric Measurement of H2O2 Production in Ehrlich Ascites Tumor Cells with the Use of CM-H2DCFDA

1. Adjust the fluorometer: Excitation – 513 nm; Emission – 530 nm; Slits (excitation and emission) ~ 3. 2. Fill the fluorometer cuvette with 3 ml of the measurement medium (NaCl 135 mM, KCl 5 mM, KH2PO4 0.4 mM, MgSO4 1 mM, HEPES 20 mM, adjusted to pH 7.4 with NaOH. Glucose 1 g/L and 1 mM CaCl2) containing 5 × 106 of EAT cells. 3. Start measurement.

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4. Add 10 mM of CM-H2DCFDA and record changes in fluorescence. 5. Make similar traces in the presence of the following additions: (a) Second trace: 2 mM oligomycin. (b) Third trace: 2 mM antimycin A. (c) Fourth trace: 2 mM FCCP. 6. An example of the results obtained is shown in Fig. 3c. 3.11. Confocal Measurements of Mitochondrial Membrane Potential in HeLa Cells

1. Plate cells on 25 mm coverslips 2 days before the experiment, in a number determined for each cell type, in order to obtain not more than 90% confluent culture on the day of the experiment. Before plating cells, the coverslips must be sterilized by UV exposure (30 J) or by temperature (200°C for 2 h). 2. Prepare a 10 nM solution of TMRM in KRB saline supplemented with glucose (1 g/L) and 1 mM CaCl2 just before loading the cells. The total volume should be calculated considering 1 ml for each coverslip and 100 ml for each drug or chemical addition executed during experiment. 3. Prepare a 10× solution of each compound required for the experiment with an exceeding volume of the TMRM solution (see Notes 12–14). 4. Wash cells twice in order to remove dead cells and cell debris, add 1 ml of TMRM solution (room temperature) to the coverslip then incubate for 20–40 min at 37°C. The correct loading time with TMRM may vary for different cell types. 5. After loading, the coverslip should be mounted in a metal cage, or a different appropriate support depending on the microscope model, and covered with 1 ml of the same TMRM solution as used for loading. 6. The coverslip and its support should be placed on the inverted confocal microscope equipped with a thermostated stage set at 37°C. Correct visualization of mitochondria should be performed using a 40–100× oil immersion objective. Optimal illumination is obtained using a 543 nm HeNe gas laser or a 561 nm solid state laser, while emission should be selected using a long pass 580 filter (see Note 15). 7. We suggest time laps with a delay of at least 10 s between each measurement step in order to avoid phototoxicity. To investigate the effect of the compound of interest (e.g., oligomycin), addition of 100 ml of the 10× concentrated solution is recommended. This is required in order to obtain a fast diffusion of the substance in the chamber. To obtain the basal fluorescence intensity level, terminate each experiment by adding 500 nM of FCCP.

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8. After the experiment, fluorescence intensity can be measured in selected regions drawn around mitochondria. An example of the results obtained is shown in Fig. 4a. 3.12. Confocal Measurement of Hydrogen Peroxide Production in HeLa Cells

1. Plate cells on 25 mm coverslips 2 days before the experiment, in a number determined for each cell type, in order to obtain not more than 90% confluent culture on the day of the experiment. Before plating cells, the coverslips must be sterilized by UV exposure (30 J) or by temperature (200°C for 2 h). 2. Prepare a 5 mM solution of CM-H2DCFDA in KRB saline supplemented with glucose (1 g/L) and 1 mM CaCl2 just before loading the cells. The total volume should be calculated considering 1 ml for each coverslip (see Note 16). 3. Prepare a 10× solution of each compound used for the stimulation or inhibition of H2O2 production in complete KRB saline. 4. Wash cells twice in order to remove dead cells and cell debris, add 1 ml of the H2DCFDA solution (room temperature) to the coverslip and incubate for 10 min at 37°C. 5. After loading, the coverslip should be mounted in a metal cage, or a different appropriate support depending on the microscope model, and covered with 1 ml of the same H2DCFDA solution used for loading. 6. The coverslip and its support should be placed on the inverted confocal microscope equipped with a thermostated stage set at 37°C. Images should be recorded with a 40–100× oil immersion objective, illuminating with 488 Argon or solid state laser. Emitted light will be preferably selected with a 505–550 band pass filter (see Note 17). 7. We suggest time laps with a delay of at least 15–30 s to avoid photoactivation not related to H2O2 production. Stimulation with chemical is obtained by adding 100 ml of the 10× concentrated solution is recommended. This is required in order to obtain a fast diffusion of the substance in the chamber. 8. After the experiment, fluorescence intensity will be measured drawing small region around each cell without touching edges to avoid artifacts (see Note 18). An example of the results obtained is shown in Fig. 4b.

3.13. Confocal Measurement of Superoxide Production in HeLa Cell

1. Plate cells on 25 mm coverslips 2 days before the experiment, in a number determined for each cell type, in order to obtain not more than 90% confluent culture on the day of the experiment. Before plating cells, the coverslips must be sterilized by UV exposure (30 J) or by temperature (200°C for 2 h).

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2. Apply the selected treatment to the sample before loading with MitoSOX Red. In the presented example, cells were incubated with 5 mM oligomycin for 15 min. 3. Prepare a 5 mM solution of MitoSOX Red in KRB saline supplemented with glucose (1 g/L) and 1 mM CaCl2 just before loading the cells. The total volume should be calculated considering 1 ml for each coverslip and should be protected from light and high temperature. 4. Wash cells twice in order to remove dead cells and cell debris, add 1 ml of MitoSOX Red solution (room temperature) to the coverslip and incubate for 15 min at 37°C (see Note 19). 5. After loading, the coverslips should be washed three times with complete KRB saline and mounted in a metal cage or in a different appropriated support depending on the microscope model. 6. The coverslip and its support should be placed on the inverted confocal microscope. Recorded images with a 40–100× oil immersion objective, illuminating with 514 Argon or 488 solid state laser. Emitted light should be selected with a 580 nm longpass filter. A thermostated stage is optional (see Note 20). 7. In our experience, MitSOX Red is not able to perform kinetics of superoxide production. After images collection, mean fluorescence intensity should be measured by drawing small regions around the bright objects for each cell. Cytosolic area should be excluded to avoid artifacts. An example of the results obtained is shown in Fig. 5. 3.14. Simultaneous Measurement of the Mitochondrial Membrane Potential and Cytosolic Calcium in HeLa Cells Using Confocal Microscope

1. Plate cells on 25 mm coverslips 2 days before the experiment, in a number determined for each cell type, in order to obtain not more than 90% confluent culture on the day of the experiment. 2. Prepare a KRB saline supplemented with 1 mM Fluo-3 AM, 10 nM TMRM, glucose (1 g/L). The total volume should be calculated considering 1 ml for each coverslip and should be protected from light and high temperature. 3. Just before loading the cells, add CaCl2 with the final 1 mM concentration to the KRB saline prepared in the previous step. 4. Wash cells twice in order to remove dead cells and cell debris, add 1 ml of room temperature KRB solution containing 1 mM Fluo-3 AM and 10 nM TMRM to the coverslip and incubate for 30 min, 37°C. 5. After loading, the coverslip should be washed with KRB/Ca2+, mounted in a metal cage or in a different appropriate support depending on the microscope model, and covered with 1 ml of the KRB saline supplemented with glucose (1 g/L) and 1 mM CaCl2.

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Fig. 5. Effect of oligomycin on mitochondrial superoxide production in HeLa cells. The mitochondrial superoxide production was measured with the use of MitoSOX Red in a confocal microscope (Subheading 3.13). Addition of oligomycin (right panel ) to the HeLa cells augmented mitochondrial superoxide production represented by increased fluorescence. These data are in line with the previous observations that in intact cells, the hyperpolarization of the inner mitochondrial membrane accelerates ROS formation.

6. The coverslip and its support should be placed on the inverted Zeiss LSM510 Confocal Microscope equipped with a thermostated stage set at 37°C. 7. Start recording the images sequentially with a 40–100× oil immersion objective, illuminating with 488 Argon or solid state laser for Fluo-3 illumination and 543 HeNe or 561 solid state laser for TMRM. Collect emitted light for the Fluo-3 in the range ³505–³535 nm and for the TMRM as total emission ³570 nm. 8. Stimulate the cells by the addition of 100 ml of the 1 mM histamine solution. 9. After the experiment, analyze the changes in fluorescence intensity with Zeiss LSM510 software. An example of the results obtained is shown in Fig. 6. 3.15. Simultaneous Measurement of the Mitochondrial Membrane Potential and H2O2 Production in HeLa Cells Using Confocal Microscope

1. Plate cells on 25 mm coverslips 2 days before the experiment, in a number determined for each cell type, in order to obtain not more than 90% confluent culture on the day of the experiment. 2. Prepare the KRB saline supplemented with 5 mM CM-H2 DCFDA, 10 nM TMRM, glucose (1 g/L), and 1 mM CaCl2. The total volume should be calculated considering 1 ml for each coverslip and should be protected from light and high temperature.

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3. Wash cells twice in order to remove dead cells and cell debris, add 1 ml of the KRB solution (room temperature) containing 5 mM CM-H2DCFDA and 10 nM TMRM to the coverslip and incubate for 20 min, 37°C. 4. After loading, the coverslip should be washed with KRB/Ca2+, mounted in a metal cage or in a different appropriate support depending on the microscope model, and covered with 1 ml of the KRB saline supplemented with glucose (1 g/L) and 1 mM CaCl2. 5. The coverslip and its support should be placed on the inverted Zeiss LSM510 Confocal Microscope equipped with a thermostated stage set at 37°C. 6. Start recording the images sequentially with a 40–100× oil immersion objective, illuminating with 488 Argon or solid state laser for CM-H2DCFDA illumination and 543 HeNe or 561 solid state laser for TMRM. Collect the emitted light for the CM-H2DCFDA in the range ³505–³535 nm and for the TMRM as total emission ³570 nm. 7. Stimulate the cells by adding 100 ml of the 1 mM histamine solution. 8. After a few minutes, when the rate of free radical production is constant, add 50 ml of 12 mM FCCP. 9. After the experiment, analyze the changes in fluorescence intensity with Zeiss LSM510 software. An example of the results obtained is shown in Fig. 7. 3.16. Measurement of Mitochondrial Respiratory Chain Activity in Human Fibroblasts with the Use of Resazurin

1. Prepare a 6 mM resazurin solution in KRB.

3.17. Measurement of Hydrogen Peroxide Production in Human Fibroblasts with the Use of CM-H2DCFDA

1. Prepare a 2 mM CM-H2DCFDA solution in KRB.

2. Remove the culture medium and wash the cells gently twice with warm KRB (see Note 6). 3. Add 0.5 ml of the resazurin solution to each well of the 24-well plate. 4. Immediately after addition, start the measurement in the kinetic mode at 510 nm excitation and 595 nm emission wavelengths (see Note 21). An example of the results obtained is shown in Fig. 8b.

2. Remove the culture medium and wash the cells gently twice with warm KRB (see Note 6). 3. Add 0.5 ml of the CM-H2DCFDA solution to each well of the 24-well plate. Immediately after addition, start the measurement in the kinetic mode at 495 nm excitation and 520 nm emission wavelengths. An example of the results obtained is shown in Fig. 8e.

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3.18. Measurement of Protein Concentration in the Plate Wells After Measurement of ROS and Mitochondrial Respiratory Chain Activity in Human Fibroblasts

1. Usually cells are plated and grow in equal density at each well; however, addition of different chemical compounds may induce cell death. Thus, protein concentration on each well should be determined after measurement to calculate appropriate values of bioenergetic parameters. 2. For spectrophotometric measurement of protein concentration, cells grown on multiwell plates must be suspended in the lysis buffer, about 500 ml per well. Protein assay is based on Bradford method. Add 2.4 ml of H2O2 to the 3 ml spectrophotometer cuvette. Depending on the cell density, add the amount of sample (that the absorbance should be between 100 and 600 spectrophotometric units) from each well to separate cuvette. Then add 600 ml of room temperature BioRad Protein Assay and shake the sample. Measure the absorbance at 595 nm(8).

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Fig. 8. Relation between mitochondrial membrane potential, respiratory chain activity, and ROS production in primary culture of fibroblasts from a healthy individual and a child with a mitochondrial disorder. C – control fibroblasts, P – patient fibroblasts. The patient used in these studies demonstrated a clinical phenotype of OXPHOS abnormality (mitochondrial encephalopathy) in muscle biopsies and in fibroblast culture. (a) DY measured with the use of JC-1 in a multiwell Plate Reader (Subheading 3.5); (b) Respiratory chain activity measured with the use of resazurin in a multiwell Plate Reader (Subheading 3.16); (c) Cytosolic superoxide production measured with the use of DHE in a multiwell Plate Reader (Subheading 3.7); (d) Mitochondrial superoxide production measured with the use of MitoSOX Red in a multiwell Plate Reader (Subheading 3.6); (e) H2O2 production measured with the use of CM-H2DCFDA in a multiwell Plate Reader (Subheading 3.17). Dysfunction of the respiratory chain in the patients’ fibroblasts was represented as a decreased respiratory chain activity and lower mitochondrial potential compared to the healthy fibroblasts. A defect in the mitochondrial respiratory chain results in higher cytosolic (c) and mitochondrial (d) superoxide production. In such cells, the rate of H2O2 production is also increased (e).

4. Notes 1. To isolate intact mitochondria, it is necessary to use lowcalcium sucrose. 2. The medium can be prepared in advance and stored in 4°C for approximately 2 weeks. 3. All solutions should be at 4°C and all equipment precooled. 4. Extreme care should be taken to avoid contamination with the ice and tap water. 5. For the assay, the reaction buffer should have room temperature. 6. Fibroblasts, plated with equal density, are grown on 24-well plates until they reach confluence, in conditions of 5% (v/v)

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CO2 in air at 37°C. The medium is changed every 2 days including the day before experiment. 7. Avoid higher JC-1 concentration as its precipitates are hard to wash out. 8. Avoid light exposure of the fluorescent probe solution and the loaded cells. 9. The time of incubation should not be longer than 10 min. 10. Keep in mind that antimycin A and oligomycin must be present during incubation with the fluorescent probe and during the measurement. 11. For the final calculation of the measured parameter, the background (basal fluorescence) should be subtracted. 12. TMRM solution may be kept for a few hours at 4°C; however, it is preferable not to add cold solution to the cells. Any changes of temperature during experiments should be avoided because it can cause alterations in TMRM distribution independently to DY. 13. In all experiments, TMRM concentration must always be the same in order to avoid differences in the redistribution of the dye in the cell. 14. Correct TMRM loading can be confirmed by a short time acquisition (i.e., 10 min). If stable fluorescence intensity is recorded, the cells are loaded correctly. If the signal is increasing during acquisition, the loading time should be increased. 15. The laser power should be kept low in order to avoid photoactivation and bleaching of the dye. With a HeNe laser, it is recommended to set the transmission lower than 10%, while with the solid state laser the power should not be higher than 30%. 16. CM-H2DCFDA solution should be protected from light and high temperature. 17. The laser power should be kept low in order to avoid photoactivation and bleaching of the dye. With a Neon laser, it is recommended to set the transmission lower than 5%, while with the solid state laser the power should not be higher than 20%. A short recording can be executed before the actual experiment at basal conditions. If fast increase in fluorescence is observed probably, photoactivation of the CM-H2DCFDA occurs. In this case, it is necessary to reduce the laser power and/or the number of the readings of the same frame (typical parameter for laser scanning confocal microscopes) and/or increase reading speed. 18. If a fast confocal system is available (i.e., spinning disk or swept field) Z stack acquisition is recommended. In conventional laser scanning confocal microscopy, Z stack acquisition will be too slow with the risk of recording artifacts.

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19. No kinetic is recorded with MitoSOX Red dye, so loading time and washing must be carefully respected in order to avoid experimental artifacts. 20. Optimal excitation wavelength declared by the manufacturer is 510 nm, but we observed that even with the 488 nm solid state laser a good signal is recordable. In this case, short exposure time is essential to avoid phototoxic stress and photoactivation. 21. For the background fluorescence subtraction, selected wells on the plate can be incubated with 1 ml of 1 M KCN stock solution before (about 15 min) and during measurement.

5. Ethics The studies with the use of human fibroblasts were carried out in accordance with the Declaration of Helsinki of the World Medical Association and were approved by the Committee of Bioethics at the Children’s Memorial Health Institute. Informed consent was obtained from the parents before any biopsy or molecular analysis was performed.

Acknowledgments This work was supported by the Polish Ministry of Science and Higher Education grants N301 092 32/3407 and NN407 075 137 for M.R.W., J.D., M.L., and J.S. J.S. was also supported by a PhD fellowship from The Foundation for Polish Science (FNP), UE, European Regional Development Fund and Operational Programme “Innovative economy.” ML is recipient of a fellowship from the Foundation for Polish Science (Program Start) and the L’Oreal fellowship (For Women in Science). P.P. and M.B. are supported by: AIRC, Telethon (GGP09128), local funds from the University of Ferrara, the PRRIITT program of the Emilia Romagna Region, the Italian Multiple Sclerosis Foundation (FISM Cod.2008/R/18), the Italian Ministry of Education, University and Research and the Italian Ministry of Health. References 1. Turrens JF (2003) Mitochondrial formation of reactive oxygen species. J Physiol 552:335–344 2. Korshunov SS, Skulachev VP, Starkov AA (1997) High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria. FEBS Lett 416:15–18

3. Miwa S, Brand MD (2003) Mitochondrial matrix reactive oxygen species production is very sensitive to mild uncoupling. Biochem Soc Trans 31(6):1300–1301 4. Wojtczak L, Teplova VV, Bogucka K, Czyz A, Makowska A, Wiechowski MR, Muszyński J,

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Evtodienko YV (1999) Effect of glucose and deoxyglucose on the redistribution of calcium in Ehrlich ascites tumour and Zajdela hepatoma cells and its consequences for mitochondrial energetics. Further arguments for the role of Ca(2+) in the mechanism of the crabtree effect. Eur J Biochem 263:495–501 5. Geromel V, Kadhom N, Cebalos-Picot I, Ouari O, Polidori A, Munnich A, Rotig A, Rustin P (2001) Superoxide-induced massive apoptosis in cultured skin fibroblasts harboring the neurogenic ataxia retinitis pigmentosa (NARP) mutation in the ATPase-6 gene of the mitochondrial DNA. Hum Mol Genet 10:1221–1228 6. Lebiedzinska M, Karkucinska-Wieckowska A, Giorgi C, Karczmarewicz E, Pronicka E, Pinton

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P, Duszyński J, Pronicki M, Wieckowski MR (2010) Oxidative stress-dependent p66Shc phosphorylation in skin fibroblasts of children with mitochondrial disorders. Biochim Biophys Acta 1797(6–7):952–960 7. Bogucka K, Teplova VV, Wojtczak L, Evtodienko YuV (1995) Inhibition by Ca2+ of the hydrolysis and the synthesis of ATP in Ehrlich ascites tumour mitochondria: relation to the crabtree effect. Biochim Biophys Acta 1228:261–266 8. Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248–254

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Chapter 13 Use of a Calcium-Sensitive Electrode for Studies on Mitochondrial Calcium Transport António J.M. Moreno and Joaquim A. Vicente Abstract Ca2+-sensitive electrode as a practical approach is used to follow Ca2+ changes in the medium and particularly useful to study mitochondrial Ca2+ uptake (or release); this method permits the continuous recording of Ca2+ movements through the mitochondrial inner membrane. In this chapter, it is described how to prepare a Ca2+-sensitive electrode, and its application on mitochondrial studies with emphasis on the mitochondrial permeability transition. Key words: Ca2+-electrode, Calcium, Isolation of liver mitochondria, Mitochondria, Mitochondrial permeability transition

1. Introduction Besides the important role in producing the cellular energy in the form of ATP by oxidative phosphorylation (»95% of the cellular ATP in the typical mammalian cell), mitochondria have been recognized to play a central role in many cellular functions where the cellular Ca2+ homeostasis is crucial (1, 2). The mitochondrion itself contributes to cellular Ca2+ steady-state since it contains a sophisticated system for sequestering and releasing Ca2+. Three modes or mechanisms of Ca2+ uptake into mitochondria were identified, the uniporter and the RAM (or “rapid mode”), although current data cannot prove clearly that the RAM is molecularly distinct from the uniporter (these two modes may represent the same protein but in different conformational forms); a third potential mechanism of Ca2+ uptake is attributed to the mitochondrial ryanodine receptor that was identified recently in heart mitochondria (3, 4).

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_13, © Springer Science+Business Media, LLC 2012

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On the other hand, mitochondria release Ca2+ via efflux mechanisms, which may be Na+-dependent or Na+-independent and, in special circumstances, the liberation of Ca2+ may be accomplish via the permeability transition pore (5–7). Mitochondrial Ca2+ uptake depends on an electrophoretic uniport movement of Ca2+ across the inner membrane in response to the electrical component (ΔYm) of the proton electrochemical potential (ΔmH+/F) established by the respiratory-chain activity. The ΔYm, amounting to 150–180 mV (internally negative), is the major contributor to the total ΔmH+/F of 200–230 mV, and the pH gradient (ΔpH) makes up the remainder (8). Consequently, respiratory activity generates large electrochemical gradients of Ca2+ across the inner membrane (9). Excessive mitochondrial Ca2+ accumulation (above a certain threshold), especially when accompanied by oxidative stress, high phosphate concentrations, and adenine nucleotide depletion, may lead to a nonselective inner mitochondrial membrane permeabilization, known as mitochondrial permeability transition (MPT). The MPT is mediated by opening the mitochondrial permeability transition pore (MPTP) that enables the passage of protons, other ions and solutes of up to 1,500 kDa (10, 11). Thus, the opening of the pore causes mitochondrial depolarization that prevents mitochondria from making ATP as long as the pore is opened. Instead, the ATP synthase is reversed to hydrolyse the glycolytic ATP leading to ATP depletion. The opening of MPTP also implicates the quick release of the previously accumulated Ca2+, the large amplitude swelling that promotes outer membrane rupture, and the release of cytochrome c in conjunction with other proapoptotic factors (12–14). MPTP has been implicated to the pathophysiology of both necrotic and apoptotic cell death (15); however, its role in triggering directly apoptosis is controversial and under great debate (16, 17). Induction of the MPTP has also been implicated in the mechanisms of cell injury caused by many chemicals (18–20). Moreover, the importance of the MPTP physiological function in cells remains unclear although some investigators have suggested that the pore may serve as a mitochondrial Ca2+ release channel (21, 22). The precise molecular identity of the membrane components of the MPTP remains controversial despite the extensive research performed in many laboratories. The adenine nucleotide translocator (ANT) is the most widely accepted candidate for the membrane components of the MPTP (15). However, some recent studies raise the possibility that mitochondrial phosphate carrier may provide the pore-forming component instead the ANT, which may only act as a regulatory component of the MPTP (23–26). The MPT can be studied in isolated mitochondria by various methods such as: (a) measuring the swelling of mitochondria

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by monitoring the associated decrease in light scattering (detected as a decrease in light absorbance with a spectrophotometer); (b) measuring the decrease of the mitochondrial electric membrane potential using fluorescent dyes or a TPP+ electrode; (c) measuring the release of accumulated Ca2+ with a Ca2+-sensitive dye (e.g., calcium green) or a Ca2+-electrode. The involvement of the MPT must be confirmed by showing inhibition of these responses by cyclosporine A. A wide range of inducers were found to activate the MPTP, such as Ca2+, phosphate, phenylarsine oxide, atractyloside, carboxyatractyloside, N-ethylmaleimide, oxidative stress, mitochondrial depolarization, while other factors promote its inhibition, as it has been observed with ADP, low pH (<7.0), bongkrekic acid, and cyclosporine A (27–29). Much of the investigation on the physiological role of mitochondrial Ca2+ transport has focused around the induction of the MPT. Since MPTP opening may be important in the pathophysiology of a wide range of diseases associated with both necrotic and apoptotic cell death, identification of new inducers or inhibitors of the MPTP might lead to development of new therapeutic agents. The MPTP opening with activators may induce cell death, which is important against malignancies, while blockers may prevent cell death which is important against degenerative and ischemia-associated pathologies. This chapter focuses on the utilization of the Ca2+-electrode and experimental procedures that can be used as a screening assay to detect novel inducers or inhibitors of the MPTP.

2. Materials Unless stated otherwise, all chemicals may be purchased from Sigma-Aldrich. 2.1. Mitochondrial Isolation from Rat Liver

1. Animals: Wistar rats, aged between 5 and 8 weeks. 2. Equipment: sharp scissors; eight 50-mL centrifuge tubes; fullsize refrigerated centrifuge (Sorvall RC-5C, Plus, SS 34 rotor, or a similar machine); vacuum pump; 60-mL Glass/Teflon motorized Potter Elvehjem homogenizer (see Note 1). 3. Solutions: Homogenization buffer comprising sucrose 250 mM, Hepes 10 mM, pH 7.4, EGTA 1 mM, defatted BSA 0.1%; washing medium comprising sucrose 250 mM, Hepes 10 mM, pH 7.2 (see Notes 2 and 3). 4. Protein assay: Biuret reagent for the determination of mitochondrial concentration.

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2.2. Construction of the Ca2+-Selective Electrode

1. Reagents: Ca2+-neutral ligand (ETH 1001, Fluka) N,N¢,-di ((11-ethoxycarbonyl)undecyl)- N , N ¢ ,4,5-tetramethyl-3,6dioxaoctane amide; o-nitrophenyl-n-octyl ether (o-NPOE) (Fluka); potassium-tetraphenylborate (TPB−) (Fluka); tetrahydrofurane (THF) (Merck); high molecular weight polyvinylchloride (PVC); CaCl2 10 mM. 2. Equipment: Five glass Petri dishes; 70-mm PVC tubing (2 mm outside diameter); scissors; scalpel; Ag/AgCl2 wire soldered to an appropriate pH meter cable.

2.3. Calibration Curve and Limit of Detection of the Ca2+ Electrode

1. Solutions: MOPS 100 mM (3-(N-morpholino)propanesulfonic acid); TAPS 100 mM (N-tris(hydroxymethyl)methyl-3-aminopropanesulfonic acid); HEPES 100 mM (N-(2-hydroxyethyl)piperazine-N¢-2ethanesulfonic acid); NTA 100 mM (nitrilotriacetic acid); HEEDTA 100 mM N-(2-hydroxyethyl)ethylenediamine N¢,N¢,N-triacetic acid; EGTA 100 mM (ethylenoglicol-bis (2-aminoethyl)-N,N,N¢,N¢-tetraacetic acid); KCl 1 M; CaCl2.2H2O 100 mM.

2.4. Measurement of Mitochondrial Ca2+ Fluxes

1. Calcium fluxes: a Ca2+ sensitive electrode sleeve; a reference electrode (we use a micro-Ag/AgCl saturated reference electrode) (Tacussel, Model MI 402); a magnetic stirrer; a circulating water bath; a thermostated open incubation chamber; a pH meter (we use a JENWAY 3520 pH meter); a suitable recorder (we use a Kipp & Zonen, BD 112). 2. Assay buffer: Sucrose 200 mM, KH2PO4 1 mM, Hepes 10 mM, adjusted to pH 7.2 by the addition of Tris base. 3. Other reagents: CaCl2 10 mM; succinate 1 M (pH 7.0); rotenone 1.5 mM (in absolute ethanol); cyclosporin A 1 mM (in absolute ethanol). All stocks are stored at −20°C.

3. Methods In this chapter, we provide methods related to the isolation of liver mitochondria and preparation of the Ca2+-electrode. We also describe assays to assess the mitochondrial Ca2+ fluxes in order to evaluate the potential of different compounds to activate or block the MPT. 3.1. Mitochondrial Isolation from Rat Liver

The following protocol is based on methods previously described (30), but see ref. 31–34 to review several mitochondrial isolation procedures. 1. Kill one rat, starved overnight to deplete the levels of endogenous fatty acids and glycogen, by cervical dislocation followed by decapitation. Let bleed from the neck under running water.

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2. Remove the liver (8–10 g) and immediately put it in a preweighed beaker, kept on ice, containing 50 mL of chilled homogenization buffer. Weigh the tissue. 3. Decant the liquid to remove any blood and add more 20–30 mL of homogenization buffer; chop the tissue with sharp scissors into the smallest possible pieces; let settle down; decant the liquid to discard the blood released; add more homogenization buffer; continue to chop; repeat the procedure several times to wash out as much blood as possible. 4. Transfer the liver to a 60-mL homogenizer (kept on ice) with 50 mL of chilled homogenization buffer. 5. Homogenize the tissue with a motor-driven pestle (500 rpm). Do not try to reach the bottom of the tube on the first pass because the vacuum effect that is produced when withdrawing the pestle may cause mechanical damage to mitochondria. Instead, it is preferable to gradually advance the pestle to the bottom of the homogenizer with successively longer strokes. Four to six strokes will be sufficient to complete an effective homogenization. 6. Distribute the homogenate equally to 2 × 50 mL centrifuge tubes, adding more ice-cold homogenization buffer to dilute each aliquot to 45 mL; balance the centrifuge tubes against each other. 7. Centrifuge at 800 × g for 10 min at 4°C. This spin sediment nuclei, large cells, tissue fragments, and red blood cells. 8. After centrifugation, remove any traces of white material (that consists of lipids) on the top of the tubes by using a Pasteur pipette and a vacuum pump. 9. Decant the supernatants into clean cold 2 × 50 mL centrifuge tubs and centrifuge at 10,000 × g for 10 min at 4°C. Let some supernatant inside the tubes to avoid disturbance of the nucleus/cell debris pellet. It is preferable to obtain a purer fraction albeit with a lower yield of mitochondria. 10. After centrifugation, decant the supernatants leaving a small volume of it in each tube. Gently swirl the liquid around the pellets to dislodge the upper fluffy layer of the mitochondrial pellet, which contain broken mitochondria; remove the remaining supernatant by using the Pasteur pipette. 11. Add a small amount (»0.5 mL) of cold washing buffer to the centrifuge tubes and very gently dislodge and resuspend the mitochondrial pellet. Let the lower part of the mitochondrial pellet (with the appearance of a white and red layer) sticked to the bottom of the tubes; this layer is composed of residual debris and red blood cells.

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12. Transfer the resuspended mitochondria to clean cold centrifuge tubes (with a few milliliters of cold washing medium in the bottom) using a variable automatic pipette (1 mL); fill to capacity and centrifuge at 10,000 × g for 10 min at 4°C. 13. Repeat the washing procedure of mitochondria. 14. Resuspend and combine the final mitochondrial pellets in 2–2.5 mL of washing buffer; the mitochondrial suspension will remain active and coupled for approximately 6 h if kept cold (keep them on ice at all times). 15. Measure the mitochondrial protein concentration using the biuret assay (see Note 4). This preparation normally yields 2.5 mL of 45 mg/mL mitochondria. 16. Before the experiments, it is important always measure the coupling of the mitochondrial preparation using an oxygen electrode. 3.2. Preparation of the Ca2+-Sensitive Electrode

1. Dissolve 3.33 g PVC in 33.3 mL THF in a 100-mL glass Erlenmeyer flask under stirring (use a fume hood). 2. Dissolve 20 mg potassium-tetraphenylborate (TPB−) in 1 mL THF in a 10 mL glass tube and combine with the previous PVC solution. Continue mixing. 3. Add 4.16 mL o-nitrophenyl-n-octyl ether (o-NPOE) and continue mixing. 4. Add the neutral carrier ETH 1001 (50 mg) and continue mixing. 5. Switched-off the fume hood and divide the mixture equally into five Petri dishes (10-cm diameter). Cover the Petri dishes with glass beakers. 6. Leave for 24–48 h to evaporate the tetrahydrofuran at room temperature. A colorless membrane of about 0.2-mm thickness is obtained. 7. Prepare electrode sleeves from a PVC tubing with an inner diameter of 2-mm and approximately 7-cm length (see Note 5). 8. Cut out a round piece of membrane, having approximately the same diameter as the PVC tubing, from the parent membrane (see Note 6). 9. Seal the round piece of membrane to the PVC tubing in the following way: pipette a droplet of THF (approximately 25 mL) to a very clean glass plate; drag on it the end of the tubing to which the membrane will be sealed; when you feel that the tubing sticks to the glass immediately place it squarely on the round piece of membrane; lightly press to promote sealing and let evaporate the THF; leave the tubing for 24 h to completely dry out; trim away any excess membrane.

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10. Check for any leaks by light sucking and blowing into the tubing with its membrane end immersed in water. 11. Fill the electrode sleeve with CaCl2 (10 mM) up to approximately 2 cm from above; the electrode should be immersed in CaCl2 (10 mM) for at least 24 h before use. 12. To complete the electrode, introduce an Ag/AgCl2 wire, soldered to an appropriate pH meter cable, in the CaCl2 10 mM inner reference solution (see Note 7). 3.3. Calibration Curve and Limit of Detection of the Ca2+-Selective Electrode

An important characteristic of an ion-selective electrode is the membrane potential, which is proportional to the logarithm of the chemical activity of the ion in solution, and varies according to Nernst equation: EM = (RT / zi F ) log (ao / ai ) where E M is the membrane potential; R is the universal gas constant; T is the absolute temperature in Kelvin; zi is the ion charge; F is the Faraday number; ao is the ion activity in the buffer; and ai is the ion activity in the inner electrode solution. For an ideal membrane, the dependence of voltage on the logarithm of ion concentration is described by a linear regression equation: F ( x) = b0 + b1 ´ x where the coefficient b1 reflects the Nernst potential of the membrane when (ao/ai) = 10 (voltage per decade of ion concentration) (slope). The ideal slope for a “good” Ca2+-electrode is 29.5 mV (at 25°C). In our experience, the Ca2+-electrodes produced as described, usually show a linear response in the range 10−7–10−3 M and a slope of 24–25 mV. 1. Prepare 50 mL of Ca2+ buffers following Table 1 (the data were taken from ref. 35). The conditions for the calibration curve and limit of detection are summarized in Table 2. 2. Determine the slope of the calibration curve and the limit of detection of the Ca2+-electrode. The calibration curve is the plot of the potential (efm) of Ca2+-selective electrode vs. the logarithm of the concentration of Ca2+.

3.4. Mitochondrial Ca2+ Transport Measurements: Effects of Potential Inducers or Inhibitors of MPTP

For a typical experiment, assemble the Ca2+-electrode in 1 mL assay buffer at 25°C. Add mitochondria (1 mg) and rotenone (2 mM) to inhibit respiratory complex I. The endogenous Ca2+ will be released from the deenergized mitochondria. Calibrate the Ca2+-electrode only after adding the compounds under study (add sequential additions of different amounts of Ca2+ in accordance with the objectives of the experiments). For example, in studying the effect of an inhibitor of the MPTP, it should be added a larger amount of

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Table 1 Preparation of 50 mL of Ca2+ buffers pCa

4 (mL)

5 (mL)

6 (mL)

7 (mL)

8 (mL)

• (mL)

MOPS 100 mM

–

–

–

5.00

–

–

TAPS 100 mM

–

5.00

–

–

–

–

HEPES 100 mM

5.00

–

5.00

–

5.00

5.00

NTA 100 mM

5.00

5.00

–

–

–

–

HEEDTA 100 mM

–

–

5.00

–

–

–

EGTA 100 mM

–

–

–

5.00

5.00

5.00

KCl 1 M

6.25

6.25

6.25

6.25

6.25

6.25

CaCl2·2H2O 100 mM

2.50

2.50

2.50

2.50

2.50

2.50

mV Brought to the specific pH value by titrating with KOH

Table 2 Composition of routine calibrating solutions with the respective pCa values (pCa = −log(free Ca2+/M)) pCa

3

4

5

6

7

8

•

pH

7.30

7.39

8.42

7.70

7.29

7.80

7.80

pH-buffer 10 mM

MOPS

HEPES

TAPS

HEPES

MOPS

HEPES

HEPES

Ca2+-ligand 10 mM

–

NTA

NTA

HEEDTA

EGTA

EGTA

EGTA

CaCl2·2H2O (mM)

1

5

5

5

5

5

–

KCl (mM)

125

125

125

125

125

125

125

Ca2+ so that in the absence of the compound the MPT could be easily observable (mitochondria release the previously accumulated Ca2+ quickly after energization as shown in Fig. 1 – panel b). Energize the mitochondria by adding succinate (5 mM). Energization of mitochondria is followed by a fast Ca2+ uptake and subsequent release of mitochondrial Ca2+ to the assay buffer. Conclusive evidence for the involvement of the MPTP can be obtained by studying the Ca2+ fluxes in the presence of cyclosporine A (1 mM), and the compounds under study. Mitochondrial Ca2+ release through the MPTP is inhibited by cyclosporine A (see Fig. 1).

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Fig. 1. A representative protocol for estimating the effects of potential inducers or inhibitors of the MPT. A specific Ca2+ electrode fitted to a recorder via a pH meter was used to record Ca2+ movements in extramitochondrial medium in an open glass thermostated-controlled reaction chamber at 25°C. Isolated liver mitochondria (1 mg) were incubated in 1 mL of assay medium containing sucrose 200 mM, Hepes-Tris 10 mM (pH 7.2), KH2PO4 1 mM (supplemented with 2 mM rotenone). Sequential additions of Ca2+ were made to calibrate the Ca2+ electrode (panel a – a: 5 mM, b: 5 mM, c: 40 mM, and d: 40 mM; panel b – a: 5 mM, b: 5 mM, c: 40 mM, and d: 60 mM). The reaction was started by adding 5 mM succinate. In panel a, 1 mM cyclosporine A plus 2 mM compound A (line 4 ), 1 mM compound A (line 2) and 2 mM compound A (line 3 ) were added to the assay medium after the mitochondria and before the calibration with Ca2+; line 1 represents the control. In panel b, 1 mM cyclosporine A plus 2 mM compound B (line 3 ) and 2 mM compound B (line 2 ) were added to the assay medium after the mitochondria and before the calibration with Ca2+; line 1 represents the control. Compound A (panel a) and compound B (panel b) were added prior to mitochondria and the calibration with Ca2+ to examine possible effects on the electrode response. The Ca2+ concentration in extramitochondrial medium decreased rapidly when succinate was added, due to Ca2+ uptake into mitochondria. Compound A acts as an inducer of the MPT since cyclosporine A reverts its action; compound B acts as a blocker of the MPT. These experiments are usually complemented with the recording of the fluctuations of the ΔY (e.g., TPP+ electrode) associated with Ca2+ fluxes and with turbidimetric studies that accompanied the swelling of mitochondria induced by the MPT.

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4. Notes 1. The motor may be substituted by a more economical option by using an electric screwdriver (e.g., Black & Decker type) with variable speed – use a low speed. 2. It has been reported that the use of mannitol (mannitol 225 mM plus sacarose 75 mM, instead of sacarose 250 mM) results in better coupled isolated mitochondria; in our experience the use of mannitol did not improve the quality of our mitochondrial preparation. 3. The BSA should be added just before the isolation procedure; we routinely add a defreezed aliquot of 5 mL of BSA 5%, previously stored at −20°C, to 250 mL of homezenization buffer. 4. The Biuret method for the measurement of mitochondrial concentration is accurate in the range of protein concentrations obtained from the protocol; other methods like the Bradford method can be used, but the mitochondria must be diluted in order to avoid saturation of the probe. 5. It is important to get a very clean and straight cut otherwise the sleeves will not stick to the membranes; use a scalpel to cut the 7-mm PVC tubes over a glass plate. 6. It can be used a leather punch belt in order to easily obtain circular pieces of membrane and avoid wasting membrane. 7. Usually, we solder a previously chlorinated silver wire (65-mm in length and 0.5-mm thick) to an appropriate pH meter cable.

Acknowledgments The authors are currently supported by grants from Fundação para a Ciência e a Tecnologia (FCT) research project PTDC/ QUI/68382/2006 (FCOMP-01-0124-FEDER 007441). References 1. Zorov DB, Krasnikov BF, Kuzminova AE, Vysokikh MY, Zorova LD (1997) Mitochondria revisited. Alternative functions of mitochondria. Biosci Rep 17:507–520 2. Zorov DB, Isanev NK, Plotnikov EY, Zorova LD, Stelmashook EV, Vasileva AK, Arkhangelskaya AA, Khrjapenkova TG (2007) The mitochondrion as janus bifrons. Biochemistry (Mosc) 72:1115–1126 3. Beutner G, Sharma VK, Giovannucci DR, Yule DI, Sheu SS (2001) Identification of a ryanodine

receptor in rat heart mitochondria. J Biol Chem 276:21482–21488 4. Altschafl BA, Beutner G, Sharma VK, Sheu SS, Valdivia HH (2007) The mitochondrial ryanodine receptor in rat heart: a pharmacokinetic profile. Biochim Biophys Acta 1768: 1784–1795 5. Gunter TE, Sheu S-S (2009) Characteristics and possible functions of mitochondrial Ca2+ transport mechanisms. Biochim Biophys Acta 1787:1291–1308

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22. Bernardi P, Petronilli V (1996) The permeability transition pore as a mitochondrial calcium release channel: a critical appraisal. J Bioenerg Biomembr 28:131–138 23. Ichas F, Mazat JP (1998) From cell calcium signaling to cell death: two conformational for the mitochondrial permeability transition pore. Switching from low- to high-conductance state. Biochim Biophys Acta 1366(1–2):33–50 24. Leung AWC, Halestrap AP (2008) Recent progress in elucidating the molecular mechanism of the mitochondrial permeability transition pore. Biochim Biophys Acta 1777:946–952 25. Leung AWC, Varanyuwatana P, Halestrap AP (2008) The mitochondrial phosphate carrier interacts with cyclophilin D and may play a key role in the permeability transition. J Biol Chem 283:26312–26323 26. Halestrap AP (2009) What is the mitochondrial permeability transition pore. J Mol Cell Cardiol 46:821–831 27. Basso E, Petronilli V, Forte MA, Bernardi P (2008) Phosphate is essential for the mitochondrial permeability transition pore by cyclosporine A and by cyclophilin D ablation. J Biol Chem 283:26307–26311 28. Zoratti M, Szabò I (1995) The mitochondrial permeability transition. Biochim Biophys Acta 1241:139–176 29. Broekemeier KM, Dempsey ME, Pfeiffer DR (1989) Cyclosporin A is a potent inhibitor of the inner membrane permeability transition in liver mitochondria. J Biol Chem 264:7826–7830 30. Pfeiffer DR, Gunter TE, Eliseev R, Broekemeier KM, Gunter KK (2001) Release of Ca2+ from mitochondria vis the saturable mechanisms and the permeability transition. IUBM Life 52: 205–212 31. Johnson D, Lardy H (1967) Isolation of liver and kidney mitochondria. Methods Enzymol 10:456–470 32. Rickwood D, Wilson MT, Darley-Usmar VM (1987) Isolation and characteristics of intact mitochondria. In: Darley-Usmar VM, Rickwood D, Wilson T (eds) Mitochondria: a practical approach. IRL Press, Oxford, pp 1–16 33. Lash L, Sall JM (1993) Mitochondrial isolation from liver and kidney: strategy, techniques, and criteria for purity. In: Lash LH, Jones DP (eds) Methods in toxicology, vol 2, Mitochondrial dysfunction. Academic, Massachusetts, pp 8–28 34. Christian F, Cipolat S, Scorrano L (2007) Organelle isolation: functional mitochondria from mouse liver, muscle and cultured fibroblasts. Nat Protoc 2:287–295 35. Tsien RY, Rink TJ (1981) Ca2+-Selective electrodes: a novel PVC-gelled neutral carrier mixture compared with other currently available sensors. J Neurosci Methods 4(1):73–86

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Chapter 14 Imaging Mitochondrial Calcium Signalling with Fluorescent Probes and Single or Two Photon Confocal Microscopy Sean M. Davidson and Michael R. Duchen Abstract The concentration of calcium ions in the mitochondria has a profound impact on mitochondrial function, modulating respiratory activity at physiological concentrations, while causing lethal damage during calcium overload. The “rhod” series of calcium sensitive fluorescent dyes tend to accumulate preferentially in mitochondria, although the reliability of mitochondrial calcium measurements depends critically on the partitioning of dye within the mitochondria which can vary between preparations. Methods are described to aid verification and quantification of the mitochondrial calcium concentration using single or two photon confocal microscopy and combining the imaging with another cytosolic calcium sensing dye. The method of linear unmixing to separate fluorescent signals based on either differing excitation or emission spectra is outlined and for the purposes of illustration is applied to the separation of rhod-2 signals originating from dye within the mitochondrial and nucleoli. Key words: Calcium, Mitochondria, Rhod-2, Confocal microscopy, Multiphoton microscopy

1. Introduction Ca2+ is not distributed evenly throughout the cell, but concentrated in microdomains, some of which are membrane-bound, such as the mitochondria (1). Ca2+ enters mitochondria via the Ca2+ uniporter in accordance with its electrochemical gradient and accumulates in the mitochondrial matrix (2). Ca2+ affects mitochondrial oxidative phosphorylation through a number of synergistic mechanisms including upregulation of the rate-limiting enzymes of the citric acid cycle (2), upregulation of the ATP synthase, and upregulation of the substrate transporter ARALAR which is activated by cytosolic rather than matrix Ca2+. Physiological calcium signalling increases (Ca2+)m in all cell types in which it has

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_14, © Springer Science+Business Media, LLC 2012

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been studied. For example, (Ca2+)m changes with each contraction in cardiomyocytes. Mitochondria may act as a spatial buffer in normal cellular calcium signalling (1, 3, 4). Mitochondrial calcium uptake is an important determinant of cell death particularly in the heart and the brain (4). In particular, mitochondrial calcium overload causes opening of a large nonspecific pore in the mitochondria – the mitochondrial permeability transition pore (mPTP) (4). This results in rapid depolarization of the mitochondrial membrane and cessation of mitochondrial oxidative phosphorylation. Prevention of mPTP opening can prevent cells from dying during ischaemia and reperfusion injury (5). Given its pathological importance and the importance of understanding how it is regulated, it is therefore necessary to be able to measure (Ca2+)m accurately. Direct measurement of intramitochondrial Ca2+ using electron probe X-ray microanalysis showed that, under physiological conditions, the concentration of free matrix Ca2+ in liver mitochondria in vivo is fairly low, at ~0.3 mM (6), though it should be kept in mind that up to 2,500 times more calcium may be present in a nonionized or bound form (7). The development of dyes which increase fluorescence when they bind Ca2+ has greatly simplified the measurement of intracellular Ca2+. Detection of mitochondrial Ca2+ was enabled in 1989 with the development of indicators based on rhodamine (8). A range of these dyes are available with different Ca2+binding affinities (Table 1), (see Note 21). With a Kd of 570 nM, rhod-2 is suitable for most studies. Intracellular accumulation is enhanced with the AM ester form of the dye (available from Invitrogen/Molecular Probes). Rhod-2/AM partitions in the mitochondria, but depending on the loading conditions used, there will normally be some dye present in the cytosol. With careful analysis, however, it can still be used to measure mitochondrial calcium.

Table 1 Kd (dissociation constant) values of a number of different mitochondrial Ca2+-sensitive dyes Dye

Kd

Rhod-2

570 nM

X-rhod-1

700 nM

Rhod-FF

19 μM

Rhod-5N

320 μM

X-rhod-FF

17 μM

X-rhod-5F

1.6 μM

X-rhod-5N

350 μM

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There are other potential limitations with the use of rhod-2 that one should be aware of. First, it is not ratiometric, i.e., since the emission spectrum does not change on binding to Ca2+, there is no way to control for efficiency of loading of the dye by comparing Ca2+-responsive and Ca2+ nonresponsive emitted fluorescence. Second, the reaction products of AM ester hydrolysis are acetate and formaldehyde, which may affect cellular functioning. Third, the presence of rhod-2 may affect Ca2+ buffering itself, since it binds to calcium, so the concentration of rhod-2/AM used should not be increased above that recommended here. Lastly, the dye may be sensitive to pH and may also accumulate in other cellular structure such as lysosomes (9). In many cells, it appears bright within the nucleolus, though this appears to be artefactual and there is no evidence it corresponds to Ca2+. It may be possible to remove some of this contaminating signal by computational methods such as linear unmixing as described in the method below, using for the purposes of illustration, the example of separating the fluorescent signal originating from the mitochondria from that originating from the nucleoli. Finally, the dyes are available over a range of affinities. The higher affinity dyes (rhod-2) may be better at detecting subtle increases in signal but will saturate when matrix calcium is high, while lower affinity dyes (e.g., rhod-5N) may not detect subtle changes but still reveal mitochondrial calcium dynamics when matrix (Ca2+)m is high. By being aware of these precautions and by following the steps outlined in the protocols below, it is possible to acquire accurate measurements of changes in (Ca2+)m. It is essential to be aware that the rhod dyes are good calcium indicators that tend to partition preferentially into mitochondria but almost invariably also leave significant concentrations of dye in the cytosol unless specific measures are taken to remove the cytosolic dye (e.g., dialysis with whole cell patch clamp recording, which has its own problems in relation to cell metabolism). Therefore, these dyes cannot be reliably used as specific mitochondrial calcium indicators using bulk measurements such as FACS or fluorimetry.

2. Materials 1. Rhod-2/AM (Molecular Probes/Invitrogen) is supplied in aliquots. Add 50 μl DMSO to a tube before use and store at −20°C for up to 1 month. Exactly the same principles apply to fluo-4/AM. 2. Imaging dish to contain coverslip and buffer while imaging cells. 3. Confocal microscope equipped with HeNe (543 nm or 555 nm wavelength) laser and ideally inverted optics (see Note 1).

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4. Imaging buffer (156 mM NaCl, 3 mM KCl, 2 mM MgSO4⋅7H2O, 1.25 mM K2HPO4, 2 mM CaCl2, 10 mM HEPES, 10 mM d-glucose, adjusted to pH 7.4 with NaOH). 5. Ca2+-free buffer (Imaging buffer + 5 μM A23187 (see Note 2) + 5 mM EGTA) to obtain the minimum (Rmin) fluorescence value. 6. High Ca2+ buffer (Imaging buffer + 5 μM A23187 (see Note 3) + 3 mM CaCl2) to obtain the maximum (Rmax) fluorescence value. Optional: 1. Intracellular buffer (130 mM KCl, 80 mM aspartate, 10 mM HEPES, 3 mM MgCl2⋅6H2O, 4 mM Na pyruvate, 0.5 mM EGTA, 3 mM Na2⋅ATP, 0.12 mM CaCl2. Adjust pH to 7.3 using KOH). 2. Permeabilization buffer (Intracellular buffer containing either 50 μg/ml saponin or 2.5 μM digitonin).

3. Methods Rhod-2/AM is taken up into cells and the AM ester is cleaved by cellular esterases. Various loading protocols can be used in order to increase the extent of dye loading into the mitochondria. A technique, which has been recommended by Lemasters and colleagues (9, 10), is used to load the dye at low temperature (RT, or even 4°C), which inhibits esterase activity and allows the dye to localize into mitochondria, then move the cells to 37°C where the dye is de-esterified and activated. However, many cells do not like being kept at 4°C. The optimum protocol depends on the cell type and will have to be determined empirically. If loading of dye can be achieved such that it is predominantly mitochondrial then it may be possible to image fluorescence using wide field fluorescent microscopy. If not, in order to reliably distinguish the mitochondrial signal from the cytosolic signal, it will be necessary to use a confocal microscope to obtain the images. Note that an “inverted microscope” with the objective approaching the coverslip from below is ideal so that the live cells can be imaged while bathed in the imaging buffer, but measurements can be made on an upright system using water dipping objectives. However, even after optimization, there may still be some mislocalization of rhod-2 to intracellular structures such as lysosomes (9), and nucleoli, which it may be possible to distinguish by using dye separation techniques such as linear unmixing. This involves taking a series of images of the same field over a range of excitation or emission wavelengths,

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then digitally separating the fluorescence into two (or more) components, and is described below. Various approaches may be used to verify the extent of mitochondrial loading, including: ensuring the majority of the dye remains in the mitochondria after plasma membrane permeabilization (Fig. 1d); loading cells with a cytosolic dye such as fluo-4/AM to confirm that the mitochondrial signal is distinct from that of the cytosol (Fig. 2); and addition of 1 μM FCCP either to depolarize mitochondria after a calcium signal to decrease (Ca2+)m or using pretreatment with FCCP before a stimulus that raises (Ca2+)m to limit mitochondrial calcium uptake. It is very important to bear in mind that after loading cells with rhod-2, mitochondrial localization may not be immediately obvious, as the signal will be weak at resting levels of (Ca2+)m. It may be necessary to stimulate the cells with an agonist to raise (Ca2+)m before the mitochondrial localization becomes evident (Fig. 2a (i and ii), b (i and ii)). Note also that calcium-independent variations in signal intensity through the cell – staining of lysosomes or the nucleoli, for example – disappear if the images are ratioed against a first “resting” image. This means extracting a baseline image from an image series and dividing the whole of the rest of the series by that first image (Fig. 2b). Under these conditions, any nonuniformities of labelling that do not change when calcium signals change disappear. To prevent the background between cells becoming very noisy (as here you are dividing very small numbers by other very small numbers and so there will be a lot of noise), one can multiply the image series by a binary mask. To achieve the binary mask is trivial in most imaging software – it requires selecting a threshold and setting all signals above the threshold to a value of unity and all pixels below the threshold to zero. Multiplying the image series by the mask makes all the signals below the threshold disappear as set to zero, while the image data of interest above threshold remains unchanged. In the example in Fig. 2, HL-1 cells – a cardiac-derived cell line that shows spectacular spontaneous calcium signals – were dual loaded with rhod-2/AM and fluo-4/AM as described in the protocol above. The sequence of Fig. 2a shows the raw rhod-2 images, in Fig. 2b the ratioed rhod-2 images while Fig. 2c shows the sequence of the fluo-4 signals (ratioed against the first image). Figure 2d shows plots of the intensity with time for each fluorophore from two regions of interest as marked (arrowhead and asterisk). Spontaneous cytosolic signals can be seen (i) and only when the cytosolic signals reached a high enough level did mitochondrial calcium rise. There are several important points illustrated here (1) at the start of the imaging, no mitochondrial signal is detectable, but mitochondria appear when calcium rises; (2) significant cytosolic rhod-2 signal remains but is distinguishable from the mitochondrial signal as the latter shows a distinct

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Fig. 1. Cells loaded with rhod-2/AM according to the standard protocol. Some fluorescence is visible originating from the mitochondria, though there is also some dye present throughout the cytosol and in the nucleoli (small spots within the nucleus) (a). By measuring the fluorescence on changing to the Ca2+-free buffer (b) and then to high Ca2+ buffer (c), it is possible to calculate the actual Ca2+ concentration, in this case, using Kd of 570 nM, we obtain a (Ca2+)mito of 196 nM. From (c), the relative concentration of rhod-2 in the mitochondria is determined to be 2.7-fold that in the cytosol. Finally, plasma membrane permeabilization (d) eliminates all but mitochondrial staining, allowing estimation of the proportion of dye localized to mitochondria (here, ~30%).

time course which is quite different to the cytosolic signal – which is in turn confirmed by the fluo-4 signal; (3) in the ratioed image local variations in signal disappear, highlighting only the areas in which the calcium-specific signal is changing. It is possible to convert the fluorescent values obtained from the microscope into actual Ca2+ concentration. This requires calibration of the fluorescence values obtained when the dye is saturated (i.e., in the presence of high (Ca2+)) (Fig. 1c) and background (i.e., in the presence of a chelator of Ca2+ such as EGTA) (Fig. 1b). These measurements are performed in the presence of a Ca2+ ionophore which equalizes the concentration of calcium throughout the cell with that outside the cell.

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a

i

ii *

b

i

iii *

*

ii

*

iii *

i

ii *

Relative Fluorescence intensity

d

Traces from

Fluo-4

ii

Rhod-2

2.0 1.5

i

iii

1.0 0.5

0

*

*

3.0 2.5

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200

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Time (sec)

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Traces from Relative Fluorescence intensity

c

*

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*

ii iii i

0

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Time (sec)

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Fig. 2. HL-1 cells were dual loaded with fluo-4AM and rhod-2AM (5 μM each for 20 min at room temperature) followed by washing. They were mounted on the stage of a Zeiss 700 confocal microscope and images acquired sequentially exciting at 488 and 555 nm, measuring emitted fluorescence at 505–550 nm (fluo-4) and at >570 nm (rhod-2). Both localized and global spontaneous calcium signals occur in these cells. Images extracted from the time sequence show a “resting” signal showing small local calcium signals in two cells (i), the peak global cytosolic signal (ii), and the residual mitochondrial signal after the recovery of the cytosolic signal (iii). The sequence (a) shows the raw rhod-2 data, in (b) the rhod-2 images ratioed against the resting signal and in (c) the corresponding fluo-4 images ratioed against the resting images. In (d) two plots are shown with the points indicated arrowhead and asterisk showing the divergence of the time course of the cytosolic calcium signal (green) and the mitochondrially localized calcium signal (rhod-2, red ).

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The following procedure describes the use of a Leica SP5 confocal microscope, but the procedure is similar on any similar confocal microscope. 3.1. Preparing the Cells

1. Place glass coverslips in the bottom of 6-well tissue culture dishes or plates (see Note 4). 2. Trypsinize the cells and plate them in the wells containing coverslips at a density sufficient to achieve ~70% confluence when they are imaged. 3. Return cells to tissue culture incubator for at least 24 h to allow them to attach and recover (see Note 5).

3.2. Loading Rhod-2/ AM into the Cells

1. In a 1.5-ml Eppendorf tube, add 5 μM rhod-2/AM + 5 μl 20% pluronic, then add 1 ml imaging buffer. 2. Replace the buffer on the cells with imaging buffer containing rhod-2/AM and pluronic. 3. Leave cells 30 min RT to take up the dye. 4. Place cells in tissue culture incubator for 20 min for de-esterification of dye. 5. Leave the cells 20 min RT to complete dye de-esterification (see Note 6).

3.3. Confocal Imaging of Rhod-2/AM Fluorescence

1. Transfer the coverslip containing the cells to the imaging chamber. 2. Wipe any liquid from underneath the coverslip using a tissue and place the chamber on the objective of a microscope, adding a drop of oil to the objective if necessary. 3. Using phase contrast or transmitted light, adjust the focus until the cells are clearly visible (see Note 7). 4. In the confocal microscope software, choose an appropriate default imaging option for imaging of a red fluorescent dye (see Note 8), i.e., excitation using the 543 nm line of the laser, and emitted light collected between 560 and 630 nm (or if a bandpass filter is not available, use a longpass filter of 560 nm). 5. Decrease the laser power to ~5% to avoid damaging the cells and increase the gain setting to maximum. 6. Start the continuous scan and adjust the focus until the mitochondria in the cell are clearly visible (see Note 9). 7. Open the pinhole setting in the software to approximately 3 Airey Units (AU) (see Note 10) and decrease the gain until a signal of ~50% saturated intensity is obtained (see Note 11). 8. Stop continuous scanning. 9. Increase the number of images averaged to four and/or decrease the scan speed to obtain a higher quality image.

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10. Start the scan to obtain the image of rhod-2 fluorescence in the cells. 3.4. Calibrating the Fluorescence to Calcium Concentration

Incubating the cells with a calcium ionophore will cause (Ca2+) in all cellular compartments to equalize to the concentration in the buffer. In a buffer containing no Ca2+ and EGTA to chelate all Ca2+, the minimum rhod-2 fluorescence value can be obtained (Fig. 1b). In a buffer containing a saturating concentration of Ca2+, the maximum rhod-2 fluorescence value can be obtained (Fig. 1c). Using these images and the Kd for the dye being used, it is possible to estimate the organellar concentration of calcium (Fig. 1a). However, this will necessarily be an estimate, since intracellular components, particular pH and the presence of heave metals can influence the dye response. 1. Replace the buffer on the cells with calcium-free buffer and wait 2 min (see Notes 12 and 13). 2. Image the cells using the same parameters as above (see Note 14). 3. Replace the buffer on the cells with high calcium buffer and wait 2 min. 4. Image the cells using the same parameters as above. 5. Using the analysis part of the software, draw a region of interest (ROI) containing a single mitochondrion or close group of mitochondria. Measure the average fluorescence intensity in the ROI to obtain Fmin (calcium-free image) and Fmax (high calcium image). 6. To calculate free (Ca2+)mito, use Fmin, Fmax, and the Kd for the dye being used (see Table 1), in the following formula from Grynkiewicz et al. (11):

[F - Fmin ] éëCa 2 + ùû = K d free [F - F ] max

3.5. Assessing the Extent of Dye Compartmentalization by Plasma Membrane Permeabilization

One approach to estimate the extent of dye that is compartmentalized in the mitochondria is to permeabilize the plasma membrane, resulting in the loss of cytosolic dye into the buffer (Fig. 1d) and calculate the proportion remaining. This does not account for mitochondria volume, however. An alternative approach is to determine the relative effective concentration of rhod-2 in the mitochondria compared to the cytosol by determining the ratio of rhod-2 fluorescence in the mitochondria compared to the cytosol in the presence of saturating (high) calcium. 1. Use the image of the cells in high calcium from step 4 of Subheading 3.4. 2. The relative effective concentration of rhod-2 in the mitochondria compared to the cytosol can be determined by drawing

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one ROI in the cytosol and one ROI around the perimeter of a mitochondria, and a third ROI in a region with no cell (“background”). Calculate the ratio according to: (mitochondria-background)/(cytosol-background). 3. Continuing from step 4 of Subheading 3.4, replacing the buffer on the cells with permeabilization buffer, which permeabilizes plasma membrane while leaving organelle membranes intact. 4. Start continuous scanning. 5. After several minutes (see Note 15), the cytosolic signal will rapidly disappear over 10–20 s. This will happen at approximately the same time in cells. Wait until all cells in the field are permeabilized and signal intensity is constant. 6. Image the cells using the same parameters as step 1 (“Organelle fluorescence”). 7. Add 1 mM MnCl2 + 5 μM A23187 to the buffer to quench all remaining dye fluorescence from the mitochondria and other organelles. 8. Image the cells using the same parameters as step 1 (“background”). 9. Using the ROI surrounding an entire cell, calculate the average fluorescence intensity of each image. 10. Express (“Organelle fluorescence” − “background”)/(“total fluorescence” − “background”) as a percentage. This represents the extent of dye compartmentalization (i.e., the percentage of rhod-2 in the cell localized to mitochondria). 3.6. Using Linear Unmixing of Emitted Wavelengths to Remove Unwanted Fluorescence Artefacts

There can be occasions when it is useful to improve the image by removing unwanted fluorescence originating from cellular autofluorescence or other artefacts. It is also possible to remove or separate the fluorescent signal of other added dyes even if they have very similar or largely overlapping fluorescent emission spectra (see ref. 12 for an example of separating rhod-2 from TMRM signal) (Fig. 3b). To achieve this it is necessary to have a confocal microscope equipped with a finely graded emission filter. For example, the Leica SP5 includes as standard the ability to perform a “lambda” scan – collecting fluorescence over a range of wavelengths. The Zeiss META addition confers a similar capability. The software then allows “linear unmixing” in which the signal is separated into that originating from the different dyes. The basic principle that follows is for a Zeiss 510 META confocal, and demonstrates how the nonspecific rhod-2 fluorescence from the nucleoli can be removed from the image (Fig. 4a). 1. Perform the steps outlined in Subheadings 3.1 and 3.3 to obtain a standard image of rhod-2 fluorescence.

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Rhod-2 IR excitation spectra

b Fluorescenceintensity

Fluorescence intensity

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0 700

800

900

Wavelength (nm)

1000

229

Rhod-2 emission spectra 1

Mitochondria Nucleoli

0 350

450

550

650

Wavelength (nm)

Fig. 3. The infrared excitation spectra (a) and emission spectra (b) for rhod-2 localized to the mitochondria (solid line) and nucleoli (dashed line). Despite the spectra for rhod-2 being very similar in both compartments, they are sufficiently different to allow separation of the signals when using multiphoton microscopy.

2. In lambda-mode (selected under “configuration control” button), using the 543 nm laser, set the start and end of the lambda stack at 550 nm and 650 nm, respectively, and the interval to 10.7 nm. Set the colour palette to “Range indicator” and using continuous scan, adjust the gain so that the signal does not saturate at any pixels. 3. View the image by clicking the “mean of ROIs” display method. 4. Using the ROI tools, draw a ROI exactly surrounding the rhod-2 fluorescence in a nucleolus, a second ROI around a region where the mitochondria are bright, and a third, large ROI in a region outside the cell where there is background signal. 5. Click on “Linear unmixing” and the software will separate the image into a new image with four panels. Panel one contains all the nonspecific and nucleolar fluorescence, panel two contains the mitochondrial rhod-2 signal, panel three contains the background (which should be dark), and panel four contains an overlay of the other three panels. Adjust the colours as desired (e.g., red for mitochondria, green for nonspecific) (Fig. 4c–e). 3.7. Multiphoton Confocal Imaging of Rhod-2 Fluorescence

Multiphoton confocal imaging is similar to confocal imaging except it is designed to use a tuneable laser in the infrared range of wavelengths, and excitation of the fluorophore (i.e., rhod-2 in this case) occurs only when two independent photons arrive simultaneously at the focal point. This design confers a number of advantages including lower toxicity, less photobleaching, and greater depth penetration in the case where imaging is performed in whole tissues. The microscope must be designed or adapted specifically for multiphoton imaging, usually including a nondescanned detector for greater light sensitivity. When using the nondescanned detector,

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Fig. 4. Either single photon confocal microscopy (a) or two photon confocal microscopy (b) can be used to visualize rhod-2/ AM fluorescence. The mitochondrial signal (arrowheads) can be separated from other contributing fluorescence sources such as nucleoli (arrows), by using linear unmixing of a series of images taken using 800–950 nm excitation wavelengths (c–e), or by linear unmixing of a series of images taken with 840-nm excitation wavelength and a lambda series of emission wavelengths ranging from 506 to 635 nm (f–h).

it is necessary to exclude all extraneous lights by covering the microscope in a light-proof sheet or cover, and normally by turning out the room lights as well. To image rhod-2 fluorescence in cells, load the dye as per the procedure above, but instead of Subheading 3.3, perform the following steps (described for a Zeiss LSM NLO microscope). Rhod-2 can be excited by any of a range

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of infrared excitation wavelengths (Fig. 3a), though usually 840 nm is appropriate. 1. Transfer the coverslip containing the cells to the imaging chamber. 2. Place the chamber on the objective of a microscope. 3. Using phase contrast or transmitted light, adjust the focus until the cells are clearly visible (see Note 16). 4. In the confocal microscope software, adjust the excitation wavelength to an appropriate value (e.g., 840 nm). 5. Collect emitted light using a bandpass filter such as 575–640 nm (or if a bandpass filter is not available, use a longpass filter of 560 nm). 6. Using the least power possible, adjust the gain and power to obtain a satisfactory image. 7. Start the continuous scan and adjust the focus until the mitochondria in the cell are clearly visible (see Note 17). 8. Decrease the gain until a signal of ~50% saturated intensity is obtained (see Note 18). 9. Stop continuous scanning. 10. Increase the number of images averaged to four and/or decrease the scan speed to obtain a higher quality image. 11. Start the scan to obtain the image of rhod-2 fluorescence in the cells. 3.8. Using Linear Unmixing of Excitation Wavelengths to Remove Unwanted Fluorescence Artefacts

As describe in Subheading 3.6, it is possible to separate out the fluorescence from different dyes, not only by their emitted spectra, but by their excitation spectra. Since it is possible to tune the excitation wavelength of the infrared laser, using a multiphoton microscope one can also perform linear unmixing of the images as described below for the Zeiss 510 NLO. 1. Begin by obtaining a standard image at 840 nm as described in Subheadings 3.1 to 3.3. 2. Configure the software so that it is imaging relatively fast over the entire area of the cell (<4 s per scan) and so that there are no saturated pixels. 3. Select and run the macro “XPrint” (see Note 19). Select the “Excitation lambda stack” tab. 4. Load the data table for the appropriate objective (see Note 20). 5. Set the start and end wavelengths to 550–650 nm, respectively, at 10 nm intervals. 6. Click “Start”.

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7. Using the ROI tools, draw a ROI exactly surrounding the rhod-2 fluorescence in a nucleolus, a second ROI around a region where the mitochondria are bright, and a third, large ROI in a region outside the cell where there is background signal. 8. Click on “Linear unmixing” and the software will separate the image into four panels. Panel one contains all the nonspecific and nucleolar fluorescence, panel two contains the mitochondrial rhod-2 signal, panel three contains the background (which should be dark), and panel four contains an overlay of the other three panels. Adjust the colours as desired (e.g., red for mitochondria, green for nonspecific) (Fig. 4f–h).

4. Notes 1. An inverted microscope is more convenient for imaging live cells in buffer since the objective approaches from below the coverslip, however, it may be possible to use a microscope in a standard orientation by using an appropriate imaging chamber such as a perfusion chamber in which the cells on the coverslip are placed at the top (facing in toward the buffer in the chamber). 2. Ionomycin may be used instead of A23187. 3. Ionomycin may be used instead of A23187. 4. In order to obtain images on sufficient resolution to distinguish mitochondria, it is essential to image the cells grown on a glass coverslip rather than imaging directly in the plastic tissue culture dish. An alternative is to grow cells on glass bottom tissue culture dishes (MatTek corporation, MA, USA). 5. If the cells are not well flattened, the mitochondria will tend to remain around the nucleus making them difficult to image. It may be necessary to allow a longer time in order for some cell types to spread out on the cover slip or to precoat the coverslip with 0.1% gelatin or fibronectin. 6. Over longer periods, the dye will be gradually extruded from the cytosol but remains in the mitochondria. It can be advantageous at this stage to return the cells to normal medium overnight, replacing the imaging buffer the following day, resulting in the fluorescence being close to 100% mitochondrial in origin. 7. Do not use a fluorescent lamp to focus the cells as this tends to cause oxidative damage and a progressive increase in dye fluorescence.

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8. On the Leica confocal, “texas red” is suitable. Other equivalent dyes are TMRE or Mitotracker red. 9. It is usually easiest to focus on the mitochondria spread out around the middle of the cell, rather than those clustered around the nucleus. 10. This increases the “thickness” of the optical slice being imaged, thus decreasing the likelihood that mitochondria will move up or down out of the imaging plane. 11. The presence of saturated (i.e., maximum intensity) pixels is determined by changing the colour scale to one indicating saturated pixels as blue. Approximately 50% intensity is selected to that there is “overhead” room for the signal to increase without saturating the measurement. 12. To replace the buffer without moving the imaging field, use a 2-ml syringe with a 20-cm piece of flexible narrow tubing slipped tightly over the needle to remove the buffer, then, before cells dry out, rapidly replace the buffer with a 1-ml Gilson pipette. 13. Some cells such as muscle cells may contract when calcium is increased, making analysis difficult. To prevent morphological alterations during calibration, cells can be depleted of ATP by 10-min pretreatment with 1 mM cyanide. 14. In order to make valid comparisons and calculations between the different treatments, it is essential that the imaging parameters (laser power, gain, objective, zoom, etc.) remain the same. 15. The exact time will depend upon the cell type. 16. Do not use a fluorescent lamp to focus the cells as this tends to cause oxidative damage and a progressive increase in dye fluorescence. 17. It is usually easiest to focus on the mitochondria spread out around the middle of the cell, rather than those clustered around the nucleus. 18. The presence of saturated (i.e., maximum intensity) pixels is determined by changing the colour scale to one indicating saturated pixels as blue. Approximately 50% intensity is selected to that there is “overhead” room for the signal to increase without saturating the measurement. 19. If not printed on one of the macro buttons, it may be necessary to install the macro first (refer to the Zeiss LSM software manual). 20. This data table contains a list of laser attenuations necessary to ensure that the power is the same at each laser wavelength (since power varies with wavelength), and must be calibrated to each objective normally used.

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21. The Kd of dyes in vivo may be different from these values determined in vitro, largely due to interfering interactions with cellular components (e.g., ref. 13). For example, the value of Kd for rhod-2 is 720 nM in intact cardiomyocytes (14).

Acknowledgements This work was undertaken at UCLH/UCL who received a proportion of funding from the Department of Health’s NIHR Biomedical Research Centres funding scheme. S.D. is supported by grant EAA17568 from the Medical Research Council. References 1. Davidson SM, Duchen MR (2006) Calcium microdomains and oxidative stress. Cell Calcium 40:561–574 2. Jacobson J, Duchen MR (2004) Interplay between mitochondria and cellular calcium signalling. Mol Cell Biochem 256–257:209–218 3. Davidson SM, Duchen MR (2007) Endothelial mitochondria: contributing to vascular function and disease. Circ Res 100:1128–1141 4. Duchen MR (2004) Roles of mitochondria in health and disease. Diabetes 53(Suppl 1): S96–S102 5. Hausenloy DJ, Yellon DM (2003) The mitochondrial permeability transition pore: its fundamental role in mediating cell death during ischaemia and reperfusion. J Mol Cell Cardiol 35:339–341 6. Somlyo AP, Bond M, Somlyo AV (1985) Calcium content of mitochondria and endoplasmic reticulum in liver frozen rapidly in vivo. Nature 314:622–625 7. Coll KE, Joseph SK, Corkey BE, Williamson JR (1982) Determination of the matrix free Ca2+ concentration and kinetics of Ca2+ efflux in liver and heart mitochondria. J Biol Chem 257:8696–8704 8. Minta A, Kao JP, Tsien RY (1989) Fluorescent indicators for cytosolic calcium based on rhodamine and fluorescein chromophores. J Biol Chem 264:8171–8178

9. Trollinger DR, Cascio WE, Lemasters JJ (2000) Mitochondrial calcium transients in adult rabbit cardiac myocytes: inhibition by ruthenium red and artifacts caused by lysosomal loading of Ca(2+)-indicating fluorophores. Biophys J 79:39–50 10. Trollinger DR, Cascio WE, Lemasters JJ (1997) Selective loading of Rhod 2 into mitochondria shows mitochondrial Ca2+ transients during the contractile cycle in adult rabbit cardiac myocytes. Biochem Biophys Res Commun 236:738–742 11. Grynkiewicz G, Poenie M, Tsien RY (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260:3440–3450 12. Davidson SM, Yellon D, Duchen MR (2007) Assessing mitochondrial potential, calcium, and redox state in isolated mammalian cells using confocal microscopy. Methods Mol Biol 372:421–430 13. Harkins AB, Kurebayashi N, Baylor SM (1993) Resting myoplasmic free calcium in frog skeletal muscle fibers estimated with fluo-3. Biophys J 65:865–881 14. Du C, MacGowan GA, Farkas DL, Koretsky AP (2001) Calibration of the calcium dissociation constant of Rhod(2)in the perfused mouse heart using manganese quenching. Cell Calcium 29:217–227

Chapter 15 Mitochondrial Permeability Transition Pore and Calcium Handling Renee Wong, Charles Steenbergen, and Elizabeth Murphy Abstract Opening of a large conductance channel in the inner mitochondrial membrane, known as the mitochondrial permeability transition (MPT) pore, has been shown to be a primary mediator of cell death in the heart subjected to ischemia-reperfusion injury. Inhibitors of the MPT have been shown to reduce cardiac ischemiareperfusion injury. Furthermore, most cardioprotective strategies appear to reduce ischemic cell death either by reducing the triggers for the opening of the MPT, such as reducing calcium overload or reactive oxygen species, or by more direct inhibition of the MPT. This chapter focuses on key issues in the study of the MPT and provides some methods for measuring MPT opening in isolated mitochondria. Key words: Calcium, Mitochondria, Pore opening, Swelling, Uptake

1. Introduction The ability of mitochondria to take up and release calcium (Ca2+) is well established. The response of mitochondria to the addition of Ca2+ depends on the amount of Ca2+ added and the level of matrix Ca2+. Nicholls first reported that mitochondria would buffer extramitochondrial Ca2+ at pCa2+ (the negative logarithm of free extramitochondrial Ca2+ concentration) of ~6.1 until matrix Ca2+ reached a level of ~60 nmol Ca2+/mg protein (1). He showed that when Ca2+ was added to isolated mitochondria they would accumulate extramitochondrial Ca2+ until they came to this set point and when EGTA was added to lower Ca2+ below that set point, mitochondria would release Ca2+ (via a Ca2+ efflux pathway such as Na+-Ca2+ or H+-Ca2+ exchanger) and return to the same extramitochondrial Ca2+ set point. At matrix Ca2+ levels above

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_15, © Springer Science+Business Media, LLC 2012

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~4 nmol/mg mitochondrial protein, extracellular Ca2+ buffering occurs because the Ca2+ efflux pathway is saturated and runs at a constant rate (e.g., 5 nmol/min/mg protein in rat liver mitochondria) that is independent of matrix or extramitochondrial Ca2+ (1). When Ca2+ is added outside the mitochondria, the Ca2+ uniporter accumulates Ca2+ until the extramitochondrial Ca2+ falls to a level at which the rate of uptake by the uniporter equals the constant rate of efflux. At this point (i.e., pCa2+ ~6.1 in rat liver mitochondria in the absence of Mg2+ and Na+), influx equals efflux and the extramitochondrial Ca2+ is maintained at a constant level. However, this set point only applies when the level of total mitochondrial Ca2+ is above ~4 nmol/mg mitochondrial protein (2). At levels below this, the efflux pathway is not constant (i.e., the Ca2+ efflux pathway is not at Vmax), and a different steady-state extramitochondrial Ca2+ concentration will be achieved. McCormack et al. (3, 4) and others have pointed out that during isolation, mitochondria accumulate Ca2+ and that unless care is taken, the levels of Ca2+ found in isolated mitochondria exceed those that occur in vivo. They further note that at matrix Ca2+ levels above 4 nmol/mg protein, the mitochondrial dehydrogenases are saturated. Therefore, if the mitochondrial dehydrogenases are regulated by changes in cellular and mitochondrial Ca2+, then the basal, unstimulated matrix Ca2+ level would be expected to be below a level that activates the dehydrogenases. If sufficient Ca2+ is accumulated by mitochondria, this leads to a large increase in permeability of the inner mitochondrial membrane and the loss of solutes with a molecular weight less than 1.5 kDa. This phenomenon, termed the mitochondrial permeability transition (MPT) pore, was described by Haworth and Hunter in 1979 (5–7). In an isolated mitochondria preparation, opening of the MPT leads to swelling of the mitochondria, which is commonly used as a measure of MPT opening. However, although mitochondrial swelling occurs with isolated mitochondria undergoing MPT, mitochondrial swelling does not appear to occur in situ because of the high cytosolic level of proteins and solutes. The MPT was largely a curiosity only of interest to mitochondrial experts until it was implicated as playing a role in apoptotic and necrotic cell death. Studies in the mid-1980s suggested that inhibition of the MPT with cyclosporin A would reduce cell dysfunction and death following cardiac ischemia-reperfusion (8). Indeed, over the ensuing decades, numerous studies have confirmed that inhibition of the MPT is cardioprotective and the cardioprotective effects of MPT inhibition have been shown to occur in humans (9). Although there is considerable interest in the MPT, the molecular identity of the MPT remains controversial. Inhibitors of the adenine nucleotide translocator (ANT) can both inhibit and activate the MPT. Bonkregic acid, which locks ANT in a conformation

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that faces the matrix, inhibits the MPT, whereas atractyloside which inhibits ANT by locking it in a conformation that faces the cytosol, activates the MPT. Based on these studies and others, ANT was suggested as a possible component of the MPT. It was speculated that under high matrix Ca2+ levels, ANT and the voltage-dependent anion channel (VDAC) would come together at contact sites to adopt an alternative conformation and form a large permeability channel (i.e., the MPT). However, studies in which ANT was genetically ablated showed that MPT was still present in mitochondria from the liver of these genetically altered mice (10). Similarly, studies in which VDAC was ablated showed that MPT remained in these genetically altered mice (11). In contrast, similar studies showed that loss of cyclophilin D resulted in a decreased sensitivity of MPT activation to calcium or during ischemia-reperfusion (12). These studies strongly supported the concept that cyclophilin D (the binding partner for the MPT inhibitor cyclosporin) was an important regulator of the MPT. It has recently been suggested that the phosphate carrier might be a component of the MPT, but this will require further validation (13, 14). In addition to a lack of understanding of the composition of the MPT, we have little understanding of whether it has any role in cell physiology or whether it exists only to mediate cell death. The MPT can function in subconductance states, and it has been suggested that these transient and/or subconductance openings might be important in regulating mitochondrial Ca2+ (15). The purpose of transient pore opening has not been established, but it has been speculated that it might serve to release matrix Ca2+ (16–18). It is generally thought that Ca2+ uptake into the matrix via the Ca2+ uniporter is more rapid than Ca2+ release via the mitochondria Na+-Ca2+ exchanger. Currently, there is controversy as to whether mitochondrial matrix [Ca2+] follows cytosolic Ca2+ transients or whether the change in matrix [Ca2+] reflects a more time averaged change in cytosolic [Ca2+]. Several studies (19, 20) report that mitochondrial [Ca2+] cycles on a beat-to-beat basis. Others such as Miyamae et al. (21) suggest that mitochondrial [Ca2+] is in the range of 0.1–0.2 μM, and increases to a higher steady-state as cytosolic [Ca2+] is elevated by increasing beating frequency or by addition of isoproterenol. One key issue is whether a mitochondrial release mechanism exists with sufficient time resolution to extrude Ca2+ from the mitochondria on a beatto-beat basis. Calcium uptake into the mitochondria might exceed the normal mitochondrial Ca2+ efflux mechanisms especially during conditions of elevated Ca2+ (e.g., isoproterenol stimulation). Therefore, the MPT might serve as a Ca2+-release valve (17). Experimental support for this hypothesis is sparse, likely due in part to the difficulty in measuring small differences in matrix Ca2+ in myocytes or cells in situ. Studies by Altschuld et al.

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showed that cyclosporin increased net 45Ca2+ uptake and reduced efflux from myocytes, consistent with MPT inhibition as being a mechanism for mitochondrial Ca2+ efflux. Ichas et al. (18) also reported that Ca2+-induced Ca2+ release from mitochondria was dependent on the MPT. A transient opening of the MPT has been shown to occur in cultured hepatocytes and HM1C1 cells (15). The MPT was measured in these cells by loading mitochondria with calcein-AM, using Co2+ to quench cytosolic calcein. This takes advantage of the lack of Co2+ entry into mitochondria. Over time, the calcein fluorescence in the mitochondria declined and this decline was inhibited by cyclosporin, suggesting that the release of calcein from the mitochondria was through the MPT. Mitochondrial membrane potential was also measured and did not decline over the same time period. These data suggest that the opening of the MPT and release of calcein was transient and occurred in only a small percentage of mitochondria at any time and therefore, did not result in a macroscopic change in mitochondrial membrane potential (15). These data provide strong evidence that transient MPT opening or MPT flickering occurs normally in cells without apparent detrimental consequences. During ischemia-reperfusion, the MPT is overwhelmed with high mitochondrial Ca2+ and its normal opening set point may also be altered by reactive oxygen species (ROS) generated at the start of reperfusion (22). In the setting of ischemia-reperfusion, the majority of the mitochondria in the cell open simultaneously, resulting in a cessation of ATP generation in the cell and increased generation of ROS rather than a few mitochondria opening transiently without detriment to the cell’s ability to make ATP. This scenario is consistent with the protection afforded by cyclosporin in protocols of ischemia-reperfusion. Low pH inhibits the MPT and low pH at the start of reperfusion has also been reported to reduce ischemia-reperfusion injury (23). It has also been suggested that postconditioning protects, at least in part, by maintaining a more acid intracellular pH during early reperfusion (24). It is also consistent with the reduction in ischemic injury that occurs with interventions that reduce the rise in cell Ca2+ during ischemiareperfusion. In fact, there is considerable data to suggest that preconditioning and many of the cardioprotective drugs mediate protection at least in part by reducing Ca2+ overload. For example, inhibitors of plasma membrane Na+-Ca2+ exchanger reduce ischemiareperfusion injury and they appear to do so by reducing the rise in cytosolic Ca2+ during ischemia and early reperfusion (25). Preconditioning has also been shown to reduce the rise in Ca2+ during ischemia (26).

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2. Materials 2.1. Mitochondria Swelling Assay

1. Isolated mitochondria (stock ~2 mg/mL; final concentration ~0.5 mg/mL). 2. Swelling assay buffer: 120 mM KCl, 10 mM Tris–HCl, 5 mM MOPS, 5 mM KH2PO4, pH 7.4. Store at 4°C. Bring to room temperature and stir to aerate before use. 3. 1–10 mM Calcium chloride stock solution in H2O. 4. UV–Vis 96-well plate reader.

2.2. Mitochondria Ca2+ Uptake

1. Isolated mitochondria (stock ~2 mg/mL; final concentration ~0.25 mg/mL). 2. Ca2+ uptake assay buffer (120 mM KCl, 5 mM MOPS, 5 mM KH2PO4, 5 mM glutamate, 5 mM malate, pH 7.4). Store at 4°C. Bring to room temperature and stir to aerate before use. Add glutamate and malate fresh. 3. 1–10 mM Calcium green™-5N tetrapotassium salt solution (Molecular Probes, Carlsbad, CA). Stock solution in H2O. 4. Calcium chloride solution: prepare a 1–10 mM stock in H2O. 5. Fluorescence spectrometer with stirring capabilities.

3. Methods for Measuring MPT 3.1. Mitochondria Swelling

MPT is commonly measured by following mitochondrial swelling. In the original study by Haworth and Hunter, they found that MPT opening resulted in a conformational change in isolated mitochondria which was accompanied by swelling that can be followed by measuring changes in light scattering. Thus, a common and convenient assay for the MPT is to measure changes in light scattering following the addition of Ca2+ to mitochondria. 1. Wash the mitochondrial pellet gently 2× to remove any EGTA present during isolation. 2. Add mitochondria (0.1 mg/well) to a 96-well plate. 3. Add swelling assay buffer to bring total well volume to 0.2 mL. 4. Measure absorbance at 540 nm for ~5 min at 20 s intervals or until a steady baseline is achieved. 5. Induce swelling with the addition of 0–0.25 mM calcium chloride and continue to monitor absorbance at 540 nm.

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3.2. Mitochondria Calcium Uptake

MPT is also commonly measured by following the ability of mitochondria to take up and retain Ca2+. In this assay, extramitochondrial Ca2+ levels are measured with a fluorescence dye as Ca2+ is added to mitochondria in small increments. Fluorescence increases from baseline with each addition of calcium. However, as the mitochondria take up the added Ca2+, the fluorescence decreases back to baseline until the mitochondria have taken up enough calcium for the MPT to occur (i.e., fluorescence increases rapidly). 1. Add 1 mL of Ca2+ uptake assay buffer containing 100 nM calcium green-5N to a stirred cuvette. 2. Add mitochondria (0.25 mg) to the stirred cuvette. Upon addition of the mitochondria, the Ca2+ in the buffer (there is always some Ca2+ contamination in the buffer) will decline because of Ca2+ uptake into the mitochondria. As discussed in Subheading 1, the mitochondria will reach a set point and extramitochondrial Ca2+ will stabilize (see Fig. 1). 3. Add successive increments of Ca2+ (see Fig. 1). The concentration of Ca2+ depends on the source of mitochondria and should be adjusted such that multiple Ca2+ additions can be made before the mitochondria release their Ca2+. Typically, additions of 10 μM Ca2+ (10 nmol/mL/0.25 mg/mL = 40 nmol Ca2+/mg mitochondrial protein) are a good concentration to begin with and if needed, the Ca2+ concentration must be adjusted to the mitochondria concentration. Initially, the mitochondria can

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Fig. 1. Calcium uptake in isolated rat heart mitochondria. Two different mitochondrial runs are overlayed. Successive additions of 10 μM Ca2+ are indicated by arrows. See text for details.

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sequester the initial additions of Ca2+. As the Ca2+ levels rise to trigger MPT opening, all of the mitochondrial Ca2+ stores are released and leads to saturation of the fluorescent indicator. References 1. Nicholls DG (1978) The regulation of extramitochondrial free calcium ion concentration by rat liver mitochondria. Biochem J 176:463–474 2. McCormack JG, Halestrap AP, Denton RM (1990) Role of calcium ions in regulation of mammalian intramitochondrial metabolism. Physiol Rev 70:391–425 3. McCormack JG, Denton RM (1980) Role of calcium ions in the regulation of intramitochondrial metabolism. Properties of the Ca2+sensitive dehydrogenases within intact uncoupled mitochondria from the white and brown adipose tissue of the rat. Biochem J 190:95–105 4. Denton RM, McCormack JG, Edgell NJ (1980) Role of calcium ions in the regulation of intramitochondrial metabolism. Effects of Na+, Mg2+ and ruthenium red on the Ca2+stimulated oxidation of oxoglutarate and on pyruvate dehydrogenase activity in intact rat heart mitochondria. Biochem J 190:107–117 5. Haworth RA, Hunter DR (1979) The Ca2+induced membrane transition in mitochondria. II. Nature of the Ca2+ trigger site. Arch Biochem Biophys 195:460–467 6. Hunter DR, Haworth RA (1979) The Ca2+induced membrane transition in mitochondria. I. The protective mechanisms. Arch Biochem Biophys 195:453–459 7. Hunter DR, Haworth RA (1979) The Ca2+induced membrane transition in mitochondria. III. Transitional Ca2+ release. Arch Biochem Biophys 195:468–477 8. Griffiths EJ, Halestrap AP (1993) Protection by cyclosporin A of ischemia/reperfusioninduced damage in isolated rat hearts. J Mol Cell Cardiol 25:1461–1469 9. Piot C, Croisille P, Staat P, Thibault H, Rioufol G, Mewton N (2008) Effect of cyclosporine on reperfusion injury in acute myocardial infarction. N Engl J Med 359:473–481 10. Kokoszka JE, Waymire KG, Levy SE, Sligh JE, Cai J, Jones DP (2004) The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature 427: 461–465

11. Baines CP, Kaiser RA, Sheiko T, Craigen WJ, Molkentin JD (2007) Voltage-dependent anion channels are dispensable for mitochondrialdependent cell death. Nat Cell Biol 9:550–555 12. Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA (2005) Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature 434:658–662 13. Leung AW, Halestrap AP (2008) Recent progress in elucidating the molecular mechanism of the mitochondrial permeability transition pore. Biochim Biophys Acta 1777:946–952 14. Leung AW, Varanyuwatana P, Halestrap AP (2008) The mitochondrial phosphate carrier interacts with cyclophilin D and may play a key role in the permeability transition. J Biol Chem 283:26312–26323 15. Petronilli V, Miotto G, Canton M, Brini M, Colonna R, Bernardi P (1999) Transient and long-lasting openings of the mitochondrial permeability transition pore can be monitored directly in intact cells by changes in mitochondrial calcein fluorescence. Biophys J 76:725–734 16. Gunter TE, Pfeiffer DR (1990) Mechanisms by which mitochondria transport calcium. Am J Physiol 258:C755–C786 17. Bernardi P, Petronilli V (1996) The permeability transition pore as a mitochondrial calcium release channel: a critical appraisal. J Bioenerg Biomembr 28:131–138 18. Ichas F, Jouaville LS, Mazat JP (1997) Mitochondria are excitable organelles capable of generating and conveying electrical and calcium signals. Cell 89:1145–1153 19. Robert V, Gurlini P, Tosello V, Nagai T, Miyawaki A, Di Lisa F (2001) Beat-to-beat oscillations of mitochondrial [Ca2+] in cardiac cells. EMBO J 20:4998–5007 20. Bell CJ, Bright NA, Rutter GA, Griffiths EJ (2006) ATP regulation in adult rat cardiomyocytes: time-resolved decoding of rapid mitochondrial calcium spiking imaged with targeted photoproteins. J Biol Chem 281: 28058–28067

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21. Miyamae M, Camacho SA, Weiner MW, Figueredo VM (1996) Attenuation of postischemic reperfusion injury is related to prevention of [Ca2+]m overload in rat hearts. Am J Physiol 271:H2145–H2153 22. Murphy E, Steenbergen C (2008) Mechanisms underlying acute protection from cardiac ischemia-reperfusion injury. Physiol Rev 88:581–609 23. Inserte J, Barba I, Hernando V, Abellan A, Ruiz-Meana M, Rodriguez-Sinovas A (2008) Effect of acidic reperfusion on prolongation of intracellular acidosis and myocardial salvage. Cardiovasc Res 77:782–790

24. Cohen MV, Yang XM, Downey JM (2007) The pH hypothesis of postconditioning: staccato reperfusion reintroduces oxygen and perpetuates myocardial acidosis. Circulation 115:1895–1903 25. Murphy E, Perlman M, London RE, Steenbergen C (1991) Amiloride delays the ischemia-induced rise in cytosolic free calcium. Circ Res 68:1250–1258 26. Steenbergen C, Perlman ME, London RE, Murphy E (1993) Mechanism of preconditioning. Ionic alterations. Circ Res 72: 112–125

Chapter 16 Imaging of Mitochondrial pH Using SNARF-1 Venkat K. Ramshesh and John J. Lemasters Abstract Laser scanning confocal microscopy provides the ability to image submicron sections in living cells and tissues. In conjunction with pH-indicating fluorescent probes, confocal microscopy can be used to visualize the distribution of pH inside living cells. Here, we describe a confocal microscopic technique to image intracellular pH in living cells using SNARF-1, a ratiometric pH-indicating fluorescent probe. SNARF-1 is ester-loaded into the cytosol and mitochondria of adult cardiac myocytes. Using 568-nm excitation, emitted fluorescence longer and shorter than 595-nm are imaged and then ratioed after background subtraction. Ratio values for each pixel are converted to values of pH using a standard curve (lookup table). Images of the intracellular distribution of pH show cytosolic and nuclear areas to have a pH of ~7.1, but in regions corresponding to mitochondria, pH is 8.0, giving a mitochondrial ΔpH of 0.9. During hypoxia, mitochondrial pH decreases to cytosolic values, signifying the collapse of ΔpH. These results illustrate the ability of laser scanning confocal microscopy to image the intracellular distribution of pH in living cells and to determine mitochondrial ΔpH. Key words: Confocal microscopy, Cytosol, Mitochondria, Myocytes, pH, SNARF-1

1. Introduction ATP is the source of energy for most biological reactions with mitochondria being the principal ATP generator in aerobic tissues like heart, brain, liver, and kidney. To synthesize ATP from ADP and phosphate via the mitochondrial F1F0-ATP synthase, mitochondria must generate a protonmotive force (Δp) across the inner membrane. Δp in millivolts equals ΔY − 60ΔpH, where ΔY is the mitochondrial membrane potential (negative inside) and ΔpH is the mitochondrial pH gradient (alkaline inside) (1). Δp also supports other energy-requiring reactions, such as ion transport and the NAD(P) transhydrogenase reaction. The ΔY component of Δp can

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_16, © Springer Science+Business Media, LLC 2012

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be visualized by confocal microscopy by any of several membrane permeant cationic fluorophores, such as rhodamine 123 and tetramethylrhodamine methylester that accumulate electrophoretically into polarized mitochondrial (2). Several techniques have been developed to measure average mitochondrial ΔpH in isolated mitochondria, cell suspensions, and cell cultures (1). However, the magnitude of ΔpH of individual mitochondria in single cells has been much more difficult to assess, since mitochondria are too small to measure ΔpH using microelectrodes. Here, we illustrate the use of laser scanning confocal microscopy to visualize pH of individual mitochondria in living cardiac myocytes under normal and hypoxic conditions by ratiometric imaging of SNARF-1 (3).

2. Materials 2.1. Buffers

1. Buffer A (5 mM KCI, 110 mM NaCl, 1.2 mM NaH2PO4, 28 mM NaHCO3, 30 mM glucose, 20 mM butanedione monoxime, 0.05 U/ml insulin, 250 μM adenosine, 1 mM creatine, 1 mM carnitine, 1 mM octanoic acid, 1 mM taurine, 10 U/ml penicillin, 10 μg/ml streptomycin, and 25 mM HEPES, pH 7.30) (see Note 1). 2. Buffer B (Joklik’s medium and medium 199 (1:1 mixture) supplemented with 20 mM butanedione monoxime, 1 mM creatine, 1 mM taurine, 1 mM octanoic acid, 1 mM carnitine, 0.05 U/ml insulin, 10 U/ml penicillin, and 10 μg/ml streptomycin). 3. Culture medium (Eagle’s minimum essential medium supplemented with 5% newborn-calf serum, 0.5 U/ml penicillin G potassium salt, 0.05 mg/ml streptomycin sulfate, and 0.5 μg/ml amphotericin B). 4. Krebs–Ringer–HEPES buffer (KRH) (110 mM NaCl, 5 mM KCI, 1.25 mM CaCl2, 1.0 mM Mg2SO4, 0.5 mM Na2HPO4, 0.5 mM KH2PO4, and 20 mM HEPES, pH 7.4).

3. Methods 3.1. Preparation of Cardiac Myocytes

Adult rabbit cardiac myocytes are isolated by enzymatic digestion, as described in ref. 3, and plated at a density of 15,000/cm2 on #1.5 glass coverslips coated with laminin (10 μg/cm2). Experiments are conducted 1 day after initial plating. Cell lines or other primary cells (e.g., hepatocytes) may be substituted for myocytes.

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3.2. Loading of SNARF-1

Intracellular pH is estimated with SNARF-1, a pH-sensitive fluorophore with a pKa of about 7.5. To load SNARF-1, cultured myocytes are incubated with 5 μM SNARF-1 acetoxymethyl ester (SNARF-1AM) for 45 min in culture medium at 37°C. During incubation, intracellular esterases release and trap SNARF-1 free acid in the cytoplasm. The cells are washed twice with KRH and placed on the microscope stage in KRH or other physiological medium like Buffer A or B. Unlike other ester-loaded fluorescent indicators, SNARF-1 loads well into mitochondria, although such loading may be cell specific. To promote better mitochondrial uptake, cells can be loaded with SNARF-1AM at a cooler temperature (4–12°C) for a longer time (4, 5).

3.3. Confocal Imaging of pH of Cardiac Myocytes

Confocal imaging of cells is performed using 568-nm excitation of an argon-krypton laser, which is near the absorbance maximum for the dye (3). At this excitation, SNARF fluorescence increases at >620 nm with increasing pH but remains unchanged at 585 nm (Fig. 1). Alternatively, excitation can be performed with the 543-nm line of a helium-neon laser. Emitted fluorescence is divided by a 595-nm long-pass dichroic reflector with the shorter wavelengths directed through a 585-nm (10-nm band pass) barrier filter and longer wavelengths through a 620-nm long-pass filter to separate detectors. Importantly, image oversaturation (pixels at highest gray level) and undersaturation (pixels with a zero gray level) should be kept to a minimum, and laser intensity should be kept at the lowest level possible consistent with an acceptable single-to-noise ratio (S/N). Because images are to be ratioed and background subtracted, S/N ratios higher than required for routine imaging are needed in both image channels. If necessary to improve S/N, binning or median filtering of pixels can be performed whereby each

Fig. 1. Fluorescence emission spectra of SNARF-1. Fluorescence emission spectra of SNARF-1 in KRH buffer at different pH. Excitation wavelength is 568 nm.

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Fig. 2. Principle of background subtraction and ratio imaging. See text for details.

pixel is reassigned a value equal to the average of 2 × 2 or 3 × 3 groups of pixels or the median value of the pixel and its adjacent pixels. If instrumentation permits, images should be acquired using the multitrack option where each wavelength is acquired alternately on a line-by-line basis. The intensity of fluorescence acquired at the two wavelengths must be corrected for background (background subtraction) (see Note 2) and then divided on a pixel-by-pixel basis (ratioing) (Fig. 2). The resulting ratios are converted to pH values based on an in situ pH calibration of SNARF-1 through the microscope optics (see Note 3). An example of measured pH in a myocyte before and after chemical hypoxia (see Note 4) is shown in Fig. 3. pH is estimated at 7.0–7.2 in cytosolic (e.g., subsarcolemmal areas) and nuclear regions and 8.0 in mitochondria, yielding a mitochondrial ΔpH of ~0.9. This gradient decreased to ~0.5 and 0 after 30 and 40 min of chemical hypoxia, respectively. After 42 min of hypoxia, the myocyte hypercontracted and died.

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Fig. 3. Confocal SNARF-1 ratio images of intracellular pH. A 1-day cultured cardiac myocyte was loaded with SNARF-1-AM (5 μM) for 45 min in culture medium at 37°C, and intracellular pH was measured by ratio imaging of SNARF-1 fluorescence before (baseline) and after 30, 40, and 42 min of chemical hypoxia.

4. Notes 1. All solutions should be prepared in deionized distilled water that has a resistance of 18.2 MΩ. 2. In confocal microscopy, detectors generate signals even in the absence of light. To quantify this dark signal, background images are collected by focusing the objective lens completely within the coverslip just underneath the cells using the same instrument settings as during acquisition of cell images. Average pixel intensity for each color channel of the background images is then determined and subtracted from each pixel of the fluorescence images of the cells at each of the two emission wavelengths. The resulting images are the backgroundsubstracted fluorescence images (see Fig. 2). 3. For in situ calibration, SNARF-1 loaded myocytes are incubated with 5 μM valinomycin and 10 μM nigericin in modified KRH buffer in which KCI and NaCl are replaced by their corresponding gluconate salts to minimize swelling (6). Images are then collected as extracellular pH is varied. Instrument settings should be the same. Alternatively, the fluorescence of SNARF-1 free acid (100–200 μM) in solution can be imaged through the microscope optics as pH is varied. After background subtraction, the >620-nm image channel is divided by the 585-nm channel on a pixel-by-pixel basis. Using thresholding to eliminate low pixel values of the extracellular space, a standard curve is created relating ratio values to pH. Lookup tables are then created assigning specific colors to different values of pH. 4. To simulate the ATP depletion and reductive stress of hypoxia, myocytes are exposed to 2.5 mM NaCN, an inhibitor of

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mitochondrial respiration, and 20 mM 2-deoxyglucose, an inhibitor of glycolysis. This treatment is termed as chemical hypoxia (3).

Acknowledgment This work was supported, in part, by grants 2-R01 DK37034, 1 R01 DK073336, and 1 R01 DK070195 from the National Institutes of Health. Imaging facilities were supported, in part, by NIH Center Grant 1P30 CA138313. References 1. Nicholls DG, Ferguson SJ (2002) Bioenergetics 3. Academic Press, London 2. Lemasters JJ, Ramshesh VK (2007) Imaging of mitochondrial polarization and depolarization with cationic fluorophores. Methods Cell Biol 80:283–295 3. Chacon E, Reece JM, Nieminen AL, Zahrebelski G, Herman B, Lemasters JJ (1994) Distribution of electrical potential, pH, free Ca2+, and volume inside cultured adult rabbit cardiac myocytes during chemical hypoxia: a multiparameter digitized confocal microscopic study. Biophys J 66(4):942–952 4. Nieminen AL, Saylor AK, Tesfai SA, Herman B, Lemasters JJ (1995) Contribution of the mito-

chondrial permeability transition to lethal injury after exposure of hepatocytes to t-butylhydroperoxide. Biochem J 307(Pt 1):99–106 5. Trollinger DR, Cascio WE, Lemasters JJ (1997) Selective loading of Rhod 2 into mitochondria shows mitochondrial Ca2+ transients during the contractile cycle in adult rabbit cardiac myocytes. Biochem Biophys Res Commun 236(3): 738–742 6. Kawanishi T, Nieminen AL, Herman B, Lemasters JJ (1991) Suppression of Ca2+ oscillations in cultured rat hepatocytes by chemical hypoxia. J Biol Chem 266(30): 20062–20069

Chapter 17 Redox Equivalents and Mitochondrial Bioenergetics James R. Roede, Young-Mi Go, and Dean P. Jones Abstract Mitochondrial energy metabolism depends upon high-flux and low-flux electron transfer pathways. The former provide the energy to support chemiosmotic coupling for oxidative phosphorylation. The latter provide mechanisms for signaling and control of mitochondrial functions. Few practical methods are available to measure rates of individual mitochondrial electron transfer reactions; however, a number of approaches are available to measure steady-state redox potentials (Eh) of donor/acceptor couples, and these can be used to gain insight into rate-controlling reactions as well as mitochondrial bioenergetics. Redox changes within the respiratory electron transfer pathway are quantified by optical spectroscopy and measurement of changes in autofluorescence. Low-flux pathways involving thiol/disulfide redox couples are measured by redox western blot and mass spectrometry-based redox proteomics. Together, the approaches provide the opportunity to develop integrated systems biology descriptions of mitochondrial redox signaling and control mechanisms. Key words: NADH, NADPH, NADH dehydrogenase, Ubiquinone, Cytochromes, Hydrogen peroxide, Glutathione, Thioredoxin-2, Redox western blot, Redox proteomics, Peroxiredoxin

1. Introduction Mitochondrial function depends upon oxidation–reduction (redox) processes with three areas especially relevant to contemporary mitochondrial research. The first is the use of energy from oxidation to support electrochemical coupling of oxidative phosphorylation (1). The second involves a small fraction of the O2 consumed by mitochondria which is converted to the so-called reactive oxygen species (ROS), superoxide anion radical (O2·−) and H2O2 (2), with mitochondria being selectively vulnerable to oxidative damage (3, 4). The third encompasses low-flux redox reactions which function in cell signaling and control (5). Another chapter in this volume (Wieckowski) addresses mitochondrial ROS generation, so this is included in this chapter. Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_17, © Springer Science+Business Media, LLC 2012

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The first section addresses measurement of steady-state redox potentials of the high-flux redox systems of the mitochondrial respiratory chain. These methods were developed over 50 years ago and currently provide a convenient but under-utilized approach to improve understanding of mitochondrial bioenergetics and respiratory control. As described by Britton Chance in 1957 (6), the mitochondrial electron transfer chain undergoes a “cushioning effect” in which the steady-state reduction of components changes with substrate conditions to maintain function of oxidative phosphorylation. Because the system functions in vivo under a nonequilibrium steady state, the steady-state redox potentials of the components within the chain are always relevant to accurate descriptions of respiratory function. A second section describes use of redox western blot methods to measure components of low-flux thiol/disulfide systems. These methods are relatively newly developed and provide the capability to selectively study mitochondrial function in intact cells and tissues due to the redox-sensitivities of specific mitochondrially localized proteins. Critically important enzymes within mitochondria are subject to redox regulation. Examples include NADH dehydrogenase which undergoes S-glutathionylation (7), apoptosis signal-regulating kinase-1 which is regulated by the redox state of thioredoxin-2 (8), and glutaredoxin-2 which is present as an inactive iron–sulfur complex until oxidatively activated (9). A subsequent section describes a mass spectrometry-based method to measure fractional reduction of cysteines in specific proteins (10). This is a highly versatile approach to profile redoxsensitive proteins which relies upon rapidly advancing technology. A final section provides some general comments concerning future needs for mitochondrial redox biology and briefly discusses related methods to study redox regulation by S-glutathionylation and S-nitrosylation. Combinations of immunoassay, mass spectrometry, and molecular manipulations provide means to define and discriminate these covalent modifications from other redox mechanisms in signaling and control. 1.1. Methodologic Limitations Due to Complexity of Mitochondria

Accurate quantification is essential to progress in redox biochemistry. However, several features of mitochondria have hindered accurate assessment of key reaction rates under relevant physiologic conditions. A first problem is that the tissue-specific regulation of mitochondria which occurs in vivo cannot readily be replicated in vitro. Because of the inability in in vitro studies to completely mimic pO2, pCO2, substrate supply, work load, and physiologic signaling as found in vivo, there is a possibility that key observations are misinterpreted. Furthermore, even if these conditions can be controlled, one is faced with the in vitro adaptations of cells, and limitations concerning function of individual mitochondria within populations of mitochondria and cells. Mitochondria are not uniform in biochemical

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characteristics and there is likelihood that spatial constraints within the cell further impact redox reactions at the individual mitochondrial level. Thus, the complexity of mitochondrial structure, function and regulation require that in vitro findings be validated in vivo and that “bottom-up” models of mitochondrial function be mirrored with “top-down” investigation. Despite the limitations of in vitro studies, measurements of mitochondrial function at the cellular level can be more reflective of physiologic rates than studies of isolated mitochondria. Fractionation of cells and isolation of mitochondria result in perturbation of rate-control characteristics (11). Isolated mitochondria invariably have some extent of physical damage, and even if this is considered minimal, there is an uncertainty about how to create a synthetic aqueous cytoplasm which appropriately reflects the intracellular environment. On the other hand, mitochondria which have been isolated from normal tissues contain proteins which are present at the correct in vivo level. Most tumor-derived cell lines have aberrant mitochondria, and mitochondrial characteristics often change dramatically when normal cells are grown in vitro. Thus, there is a need to study mitochondrial functions at multiple levels of organization. Importantly, the methods described in this chapter can be applied at all levels of organization. Thus, an advantage of available quantitative redox biology methods is that they can provide a foundation for a comprehensive understanding of mitochondrial respiratory control. 1.2. Steady-State Reduction Potentials as Quantitative Descriptors in Redox Biology

As described elsewhere in this volume, quantitative information for specific biochemical processes can be obtained using isolated mitochondria and submitochondrial fractions. These preparations provide mechanistic detail that cannot be obtained with more intact systems. There is a critical need to integrate the information obtained from mitochondria and submitochondrial fractions with data obtained from cellular and in vivo models. A particular challenge lies in the difficulty of knowing whether kinetic principles or thermodynamic principles govern specific redox processes. Measurement of steady-state redox potential (Eh) values provides means to characterize operational characteristics of mitochondria without specific knowledge of reaction rates. Eh values are reduction potentials for electron acceptor/donor pairs relative to a standard hydrogen electrode (12). By definition, the reduction potentials are given for an acceptor/donor pair, e.g., pyruvate/lactate. The biological and medical research literature more commonly uses the inverted expression of the donor/acceptor pair, e.g., lactate/pyruvate; while technically incorrect, common use requires that regardless of how the couple is ordered, Eh expressions be interpreted as reduction potentials, unless explicitly stated otherwise. In this chapter, we use donor/acceptor to refer to a redox couple and the term Eh with the acceptor to define the couple, e.g., for the GSH/GSSG couple,

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EhGSSG refers to the reduction potential for the reaction GSSG + 2e- + 2H + « 2GSH . For proteins, we use the protein abbreviation with the redox-sensitive thiols implied, e.g., for thioredoxin-2, EhTrx2 refers to the reduction potential for the active site dithiol/disulfide couple. In biological systems, Eh values are calculated using the Nernst equation, Eh = Eo + RT / nF ln ([acceptor ] / [donor]) , where Eo is the standard potential for the couple, R is the gas constant, T is the absolute temperature, n is the number of electrons transferred, and F is Faraday’s constant. The difference in Eh values between two couples, DEh, is related to Gibbs free energy for the electron transfer according to the equation DG = nF DEh . Changes in Dy and DpH depend upon the DEh for electron transfer, yet countless studies of Dy, proton leak, ROS production, etc., have been performed and interpreted without the knowledge of Eh values for donors or DEh for donor and acceptor couples under the conditions of assay. Because the validity of interpretation depends upon the energetics determined by the redox potentials of the respective upstream and downstream couples in the electron transfer pathway, there is a need to incorporate measurements redox potentials into bioenergetic descriptions. 1.3. Steady-State Redox Potentials of Mitochondrial Electron Transfer Components

In vivo, the mitochondrial electron transfer pathway operates under nonequilibrium steady-state conditions. The principles for measurement are derived from the historically important discovery of the mitochondrial cytochrome chain by David Keilin, in which oxidation–reduction reactions were characterized in terms of spectral changes in the cytochromes (Fig. 1) (13, 14). While the rates of electron transfer through the entire pathway are conveniently measured in terms of O2 consumption, measurement of specific donor pathways is more difficult. However, changes in steady-state potentials can be readily measured for many of the components because of their inherent spectral and fluorescent properties (Fig. 2) (15). The materials needed to perform experiments include appropriate spectrophotometry or fluorometry instrumentation, appropriate means to calibrate the system for complete oxidation and reduction and means to control O2 and substrate availability. Spectral changes are measured as changes in light absorbance by a chemical at a specific wavelength defined by Beer’s law, A = eCl, where A is absorbance, e is the extinction coefficient, C is the concentration, and l is the pathlength. e is usually given as the millimolar extinction coefficient and the pathlength is 1 cm in standard spectrophotometers. Studies of steady-state levels of oxidation of respiratory chain components in isolated mitochondria and cell suspensions can be obtained in 1 cm cuvettes or in small beakers (16) with appropriate positioning of light source and detector. With appropriate instrumentation, absorbance changes

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Fig. 1. Changes in steady-state redox levels of mitochondrial components are based upon absorbance characteristic as shown by this original study of yeast (13). The reduced pyridine nucleotides, NADH and NADPH, absorb light at 320 nm. Oxidized flavoproteins absorb light at 455 in the 450–500 nm range. Cytochromes have distinct absorbance in the reduced forms. Reproduced with permission from the publisher.

Fig. 2. Measurement of changes in steady-state levels of oxidation in the mitochondrial cytochrome chain. Changes in steady-state oxidation of (a) cytochrome a3 and (b) cytochrome b are illustrated in this original study of Chance (15). In a, the response of heart-muscle mitochondrial preparation to metabolism of added fumarate is shown. In b, changes in steady-state oxidation are observed in response to added succinate, with the cytochromes approaching complete reduction as O2 becomes exhausted. Reproduced with permission from the publisher.

can be measured in perfused organs and adherent cells (17). In all cases, light scatter can be limiting so that dual-wavelength spectrometry is used to minimize this problem (13, 17). Stock reagents needed for total oxidation are either 100 mM potassium ferricyanide (added at 5 ml/ml incubation) or 1 mM FCCP (carbonylcyanide p-trifluoromethoxyphenylhydrazone) in ethanol (added at 1 ml/ml incubation). Reagent needed for total reduction is sodium dithionite. The latter must be maintained dry

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so that only a few grains can be added; if a solution is used, this is prepared fresh as a 1-M solution so that only 1 ml/ml incubation is needed to effect complete reduction. It should be noted that reduction by dithionite is slower with larger amounts of dithionite due to altered chemical reactions. 1.4. NADH/NAD + Measurement

The NADH/NAD+ couple provides a central electron carrier connecting oxidation of many food-derived metabolites and energy production. Enzyme-coupled and HPLC assays are available to measure NADH and NAD+ concentrations (18, 19), and with selective solubilization of cell membranes, measurements of mitochondrial contents can be obtained (20). However, a substantial fraction of the total NADH + NAD+ pool is protein bound, so the measurements of the total amounts of NADH and NAD+ in the mitochondria do not allow accurate estimate of EhNAD+. Studies of different dehydrogenase reactions in rat liver and hepatocytes showed that the reaction catalyzed by b-hydroxybutyrate dehydrogenase is near equilibrium so that the b-hydroxybutyrate/ acetoacetate ratio can be used to calculate the matrix EhNAD+, which is about 318 mV (20). Because Eo for NADH/NAD+ is −337 mV, this means that free NAD+ is present at a considerably higher concentration than free NADH. This characteristic means that the energetics of Complex I can be altered by the rates of the NAD+linked dehydrogenases. Assay of b-hydroxybutyrate and acetoacetate can be performed with several standard approaches, including on a fluorescence plate reader using enzyme-coupled assays (19), by 1H-NMR spectroscopy (21) or gas chromatography–mass spectrometry (22). EhNAD+ is then estimated using the Nernst equation, the b-hydroxybutyrate and acetoacetate concentrations, an Eo value of −297 mV (20) and the assumption that the b-hydroxybutyrate dehydrogenase reaction is at equilibrium. It must be pointed out that it is not clear whether this reaction is at equilibrium in all mitochondria so this assumption will require validation for extension to other cell types and tissues. Measures of relative changes in the NADH/NAD+ couple can be readily obtained by absorbance and fluorescence methods. NADH has an absorbance at 340 nm (extinction coefficient, 6.2 mM−1 cm−1) which is not present in NAD+. Although not directly useful for Eh determination, measurement with dual-wavelength spectroscopy can provide a sensitive means to measure relative changes. In this approach, absorbance at 375 nm is used as a reference to control for light scatter so that absorbance change can be used to provide a measure of the absolute change in NADH concentration in intact cells (23). More sensitive detection of changes in NADH redox state is obtained by measuring fluorescence of NADH with excitation at 366 nm and emission at 450 nm (24, 25). Protein-binding results in enhanced NADH fluorescence efficiency, and fractionation studies have shown that most of the NADH fluorescence is associated

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with mitochondria. Consequently, relative changes in fluorescence of NADH provide means to measure changes in redox state of this couple in intact cells and tissues. This approach has been used effectively to study redox control in cell culture, often described as measurement of “autofluorescence” because it does not require addition of fluorescence probes. The flow of electrons from substrates to NAD+ is regulated at many levels to balance utilization of carbohydrate, fat, and amino acid-derived precursors. Some of the key dehydrogenases are activated by Ca2+, and several of the monocarboxylate, dicarboxylate, and tricarboxylate substrates are exchanged by antiporters. Multiple fatty acid transporters control utilization of different chain lengths, and variations in amino acid transporters also contribute to rates of utilization of these substrates for mitochondrial respiration. The relatively limited range of substrates and conditions used for most studies of isolated mitochondria raises the possibility that key regulatory mechanisms related to supply of reducing equivalents to maintain EhNAD+ in different tissues remain to be discovered. To date, relatively limited efforts have been made to mimic physiologically relevant metabolic conditions in studies with isolated mitochondria. 1.5. Coenzyme Q and Complex III

Electron transfer from NADH to ubiquinone (CoQ) occurs through Complex I, a redox-driven proton pump. This is a large protein complex containing a covalently bound, redox-active flavin, as well as multiple iron–sulfur centers. The DEh for the overall reaction is determined by the EhCoQ relative to EhNAD+. CoQ exists as three redox forms, ubiquinone, ubiquinol, and ubisemiquinone. The redox characteristics of quinones have recently been reviewed (26); CoQ is hydrophobic and mostly membranal, estimates of EhCoQ are obtained using the Nernst equation with relative amounts of ubiquinol and ubiquinone obtained by HPLC (27). Because CoQ is an intermediate between Complex I and Complex III and also substrate for Complex II and a number of other substratelinked dehydrogenases (e.g., acyl CoA dehydrogenases), studies of Complex I function are subject to variable energetics due to steadystate reduction of the CoQ pool. In this way, changes in DEh can alter the energetics for maintenance of Dy independently of leak currents. Thus, there is a need to include DEh measurements along with Dy and O2 consumption measurements. Absorbance and fluorescence of the flavin in complex III change upon reduction, and these changes have also been used to measure redox in cells and intact tissues (28). In contrast to the increases in absorbance and fluorescence of NADH upon reduction, flavoproteins have a decrease in absorbance and fluorescence upon reduction. Fluorescence is measured with excitation at 436 nm and emission at 570 nm, largely a measure of mitochondrial flavoproteins (29). Because there are multiple flavoproteins which can contribute to the signals, there is a limitation to the mechanistic

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information which can be derived. On the other hand, ratiometric studies of the NADH fluorescence and flavoprotein fluorescence provide very sensitive indicators of tissue anoxia which can be used to visualize regional differences in oxidation due to pathophysiologic changes in vivo (24, 25). With modern instrumentation, these approaches could provide improved capabilities for in vivo study of mitochondrial redox reactions. 1.6. Cytochromes

Complex III and Complex IV contain hemoproteins termed “cytochromes” which are readily detected by spectrophotometry. These were used as early as 1952 to measure mitochondrial redox changes in cells (15). The measurements of the redox states of cytochromes depend upon the absorbance characteristics of the hemes in the cytochromes, which have more intense absorbance in the reduced forms (Fig. 1) (13). In the visible range, the absorbance bands are termed a, b, and g (Soret) absorbance bands. While the Soret band has the greatest extinction coefficients, it occurs at the shortest wavelength where there is the greatest light scatter in preparations containing mitochondria; consequently, measurements in this region can be more prone to error due to this light scatter. In addition, there is considerable overlap of signals from heme b and c, and these can be further obscured by signals from myoglobin or hemoglobin (30). Consequently, measurements in the Soret region are usually limited to cytochromes a3. Cytochromes a and a3 contain heme a, which is structurally distinct from heme b, present in hemoglobin, myoglobin, catalase, and b- and c-type cytochromes. Cytochromes a and a3 also absorb light in the near infrared region, and this has provided a means to measure oxidation in vivo (17). Considerable spectral overlap occurs with the b bands, so most measurements are obtained with the a bands. Cytochromes b and c contain heme b (Fe-complexed protoporphyrin IX), but differ in absorbance because the heme is covalently bound in cytochromes c and c1 but not in bH or bL. Other minor differences in absorbance maxima occur due to the interactions with amino acids in the vicinity of the heme. Addition of ligands such as KCN, which bind to the O2 site, can be used to discriminate cytochromes a and a3. However, this is not useful for steady-state redox measurement because it blocks electron flow. Complex III accepts two electrons from CoQ and transfers one to cytochrome c1 and the other to the cytochrome bH and bL pair. The electron from cytochrome c1 is then transferred to cytochrome c. While the electron from cytochrome b is transferred back to CoQ. The system is a proton pump, with energetics of proton pumping probably defined by steady-state potential of the donor ubiquinol/ubisemiquinone couple and cytochrome c1. As above, changes in the steady-state Eh values determine the energetics available for proton pumping. Consequently, interpretations of Complex III function in generation of DmH+, ROS generation, or support of ATP production, are limited by changes in steady-state Eh values which determine the available DEh.

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Steady-state reduction of cytochromes b is measured by dual-wavelength measurement with 575 as an isosbestic point. Absorbance maxima for the two forms are at 561 and 566 nm, but the signals overlap considerably so that redox changes of the mixture are most readily obtained by measuring absorbance at 562 nm relative to 575 nm. The extinction coefficient value for 562 nm minus 575 nm is 23 mM−1 cm−1. Fractional oxidation under steadystate conditions is obtained by use of an uncoupler to approximate 100% oxidation followed by the addition of sodium dithionite to obtain 100% reduction. Because dithionite breaks down rapidly in solution, complete reduction is usually achieved by the addition of a few grains of solid sodium dithionite. FCCP and CCCP (carbonylcyanide m-chlorophenylhydrazone) are commonly used as uncouplers to obtain maximal oxidation, but this approach assumes that the only limitation to oxidation is the existence of the proton gradient. Because this assumption may not be valid, an alternative means to obtain maximal oxidation is to use a chemical oxidant such as 0.5 mM potassium ferricyanide (31). In intact organs and some cell preparations, the fully reduced state can be approximated by removal of O2 supply or addition of potassium cyanide (6, 15); however, these treatments may not result in complete reduction due to endogenous respiratory control characteristics. Cytochrome c has an absorbance maximum at 550 nm in the reduced form while cytochrome c1 has a maximum at 554 nm. The two are not easily resolved spectrally in cells or tissues; therefore, the two are measured together as cytochrome c + c1. For these measures, dual wavelength spectroscopy is used by measuring 550 nm minus 540 nm as an isosbestic point. The extinction coefficient is 19 mM−1 cm−1. This measurement is usually expressed relative to maximal oxidation and reduction as performed for cytochromes b. Steady-state reduction of cytochromes a and a3 is also frequently measured together as cytochrome a + a3, using the wavelength pair 605 and 630 nm (32). The extinction coefficient is 13.1 mM−1 cm−1. 1.7. Oxygen Consumption Rate

Changes in redox state of cytochromes can be used to measure O2 consumption rate in closed systems where known amounts of O2 are added (15, 23). Typically, the system is allowed to consume all O2 and become anoxic. Rapid addition of a known amount of O2 then results in oxidation of the cytochromes, and the time required to consume the O2 and become reduced allows calculation of O2 consumption rate. Because the rate of O2 consumption decreases when O2 concentration reaches the low micromolar range, measurements are calculated from the difference in time to consume different amounts of added O2. This approach has been directly validated relative to measurements with a Clark-type electrode (23). With this approach, the duration of anoxia prior to addition of O2 pulses affects the O2 consumption rate (33), necessitating care in the timing

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of the measurements. A variation on this approach has also been used to measure rates of catalase activity in terms of the amount of O2 produced and cytochrome oxidized following addition of H2O2 under anaerobic conditions (34). 1.8. NADPH/NADP + Measurement

As indicated above, mitochondria contain low flux redox systems that are qualitatively different from those which support oxidative phosphorylation and energy metabolism. Although originally described in terms of their role in protection against oxidative stress, accumulating evidence shows that these systems function in regulation of the redox states of cysteine (Cys) residues of proteins, controlling their structures and activities. Consequently, assays to measure fractional reduction of specific Cys residues of proteins are of considerable importance to understand mitochondrial signaling and control. Important conceptual advances in understanding redox regulation of cell functions have come from redox proteomics methods, some of which are especially useful to measure mitochondrial redox systems. These systems are oxidized by the endogenous mitochondrial generation of O2·− and H2O2. An early estimate that H2O2 production by mitochondria is about 1% of the total O2 consumption rate (2) is frequently repeated, but accurate values for mitochondria in intact systems are not available. Measurements obtained with isolated mitochondrial preparations are likely to provide exaggerated values due to the presence of damage mitochondria and the analysis under supraphysiologic conditions of oxidizable substrates and pO2. On the other hand, because there is also evidence for electron transfer bypass, in which NADH dehydrogenase generates O2·−, the O2·− is transported into the intermembrane space according to the electrochemical gradient, and the O2·− donates the electron to cytochrome c (35). The rate of this pathway is not known, so that there is a possibility that the estimated rate of 1% could also be an underestimate. Available methods to measure oxidant production by mitochondria under relevant in vivo conditions are difficult to calibrate so that better approaches are needed. Nonethe-less, the data are accurate enough to allow the conclusion that only a small fraction of the total O2 consumed by cells is converted by mitochondria to oxidants. Because thiol-dependent regulatory mechanisms are dependent upon these oxidants for oxidation, this low rate of oxidant production defines redox regulatory systems as “low-flux” systems when compared to mitochondrial respiration. The practical meaning is that minor disruption of normal high-flux electron transfer by the respiratory chain can produce oxidants at rates which are comparable to or higher than the rates which normally function in regulation. These issues have been previously reviewed and provide the basis for a definition of oxidative stress which includes the disruption of redox signaling and control pathways (36, 37).

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While rates of electron transfer through the low flux redox signaling and control pathways are not known, information about their function can be obtained from measurements of steady-state redox potentials. The thiol-dependent systems are maintained by electron transfer from NADPH. The NADPH/NADP+ couple in mitochondria is maintained at relatively negative Eh value, about −415 mV in liver (20). As with the NADH/NAD+ couple, direct measurement of total concentrations does not give a reliable measure of the potential because of extensive protein binding. Consequently, better estimates are obtained in liver by measuring the glutamate, ammonia, and 2-oxoglutarate concentrations and calculating the Eh with the assumption that the reaction is at equilibrium. Alternatively, isocitrate, 2-oxoglutarate, and CO2 can be used (20). Little information is available concerning whether these reactions are at equilibrium in different tissues, so some extent of validation would be important in use of this approach.

2. Materials 2.1. GSH/GSSG Sample Buffer and Derivatization Reagents

1. 10% PCA/BA (10% perchloric acid/0.2 M boric acid) is prepared by adding 71 ml of 70% perchloric acid, 6.2 g boric acid (0.2 M) and adjusting total volume to 500 ml. 2. 10% PCA/BA with internal standard is made by adding 1.38 mg g-glutamylglutamate (10 mM) to the solution above. 3. 20 mg/ml dansyl chloride solution prepared in 100% acetone. 4. 9.3 mg/ml iodoacetic acid solution prepared in water. 5. 1% digitonin (w/v) solution in water. 6. Chloroform. 7. Acetone.

2.2. Mobile Phase Buffers for GSH/GSSG HPLC Determination

1. Solvent A: 80% methanol in water.

2.3. SDSPolyacrylamide Gel Electrophoresis

1. Running buffer (10×): 250 mM Tris–HCl, 1.92 M glycine, 1% SDS.

2. Solvent B: 64% methanol, 4 M sodium acetate buffer, pH 4.6.

2. 30% Acrylamide/bis solution 37.5:1 (2.6% C) (BioRad, Hercules, CA) and N,N,N,N¢-tetramethyl-ethylenediamine (TEMED) (Sigma). 3. Ammonium persulfate (APS): Prepare a 10% stock solution and store at −20°C. 4. Precision plus protein standards (BioRad).

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5. Gel casting stock solutions: 1.5 M Tris–HCl (pH 8.8) to be used for casting the resolving gel; 1.0 M Tris–HCl (pH 6.8) to be used in casting the stacking gel; 10% SDS. 6. Mini Protean III cell (BioRad) and Power Pac 200 power source (BioRad). 2.4. Redox Western Blotting (Trx2, Prx3, TrxR2)

1. Towbin’s transfer buffer: 25 mM Tris–HCl, 192 mM glycine, 20% methanol. Store at 4°C. 2. Nitrocellulose membrane (BioRad) and extra thick filter paper (7.5 × 10 cm) (BioRad). 3. Trans-blot SD semidry transfer apparatus (BioRad) and Power Pac 200 power source (BioRad). 4. PBS with Tween-20 (PBS-T)/wash buffer: add 2 mL of Tween-20 (Sigma) to 1 L of 1× PBS. 5. Blocking buffer/antibody dilution buffer: 1:1 mixture of PBS-T and Li-Cor blocking buffer (Li-Cor, Lincoln, NE). 6. Primary antibody: Rabbit antisera against Trx2; Prx3 mouse monoclonal antibody (Abcam, Cambridge, MA); TrxR2 rabbit polyclonal antibody (AbFrontier, Korea). 7. Secondary antibody: goat-antirabbit or goat-antimouse Alexafluor680 antibody (Invitrogen, Eugene, OR). 8. Li-Cor Odyssey infrared imager (Li-Cor).

2.5. Cell Lysis and Protein Alkylation: Trx2 Redox Western Blot

1. 10% Trichloroacetic acid solution (Sigma). 2. 100% Acetone (Sigma). 3. Phosphate-buffered saline (PBS) (Mediatech). 4. Disposable cell scraper (Fisher). 5. Alkylation buffer: 50 mM Tris–HCl (pH 8), 0.1% SDS, 15 mM 4-acetoamido-4¢-maleimidylstilbene-2,2¢-disulfonic acid (AMS) (Invitrogen). 6. Nonreducing sample loading buffer (5×): 300 mM Tris–HCl (pH 6.8), 50% glycerol, 1% SDS, 0.05% bromophenol blue.

2.6. Cell Lysis and Protein Alkylation: Prx3 Redox Western Blot

1. DMEM/F-12 (1:1) (with L-glutamine) cell culture medium (Mediatech, Manassass VA) supplemented with 10% fetal bovine serum (Atlanta Biologicals, Atlanta, GA) and 1% penicillin/ streptomycin (Hyclone, Logan, UT). 2. Paraquat (Sigma, St. Louis, MO) 10 mM stock solution in water. Prepare stock solution and use within 15 min of preparation. 3. PBS (Mediatech, Manassass, VA). 4. Alkylation buffer: 40 mM Hepes, 50 mM NaCl, 1 mM EGTA, Protease inhibitor cocktail (Sigma), 100 mM N-ethylmaleimide (NEM). Add NEM to buffer immediately prior to use.

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5. 25% CHAPS stock solution for cell lysis. 6. Disposable cell scraper (Fisher). 7. Nonreducing sample loading buffer (5×): 300 mM Tris–HCl (pH 6.8), 50% glycerol, 1% SDS, 0.05% bromophenol blue. 2.7. BIAM Modification and Pull-Down: TrxR2 Redox Western Blot

1. Biotin iodoacetamide [N-(biotinoyl)-N¢-(iodoacetyl)ethylenediamine, BIAM, Molecular Probes]: Dissolved in 50 mM Tris buffer (pH 6.8) to make a stock solution at 0.1 M concentration. 2. Iodoacetamide (IAM, Sigma-Aldrich Co.): Dissolve in Tris buffer at 5 mM concentration. 3. Lysis buffer A [50 mM Bis–Tris–HCl (pH 6.5), 0.5% Triton X-100, 0.5% deoxycholate, 0.1% SDS, 150 mM NaCl, 1 mM EDTA, leupeptin, aprotinin, and 0.1 mM PMSF] containing 10 mM BIAM. 4. Protein G sepharose (Sigma-Aldrich, St. Louis, MO).

2.8. Cell Lysis and ICAT Reagents: Redox Proteomics

1. 10% TCA.

2.9. Mass Spectrometry-Based Redox Proteomics

1. Ultimate 3000 nanoHPLC system (Dionex) with a nanobore column (0.075 × 150 mm Pepmap C18 100 Ǻ, 3 mm, Dionex).

2. Denaturation buffer: 50 mM Tris, pH 8.5, 0.1% SDS. 3. ICAT assay kit (Applied Biosystems, Foster City, CA).

2. QSTAR XL Q-TOF mass spectrometer (Proxeon Biosystems). 3. ProteinPilot software (Applied Biosystems).

3. Methods 3.1. GSH/GSSG Measurement in Cultured Cells

3.1.1. Sample Derivatization

Mitochondria contain two central systems controlling protein thiol/disulfide states, one dependent upon GSH and the other dependent upon a mitochondria-specific thioredoxin, thioredoxin-2 (Trx2). These systems are parallel, nonredundant systems (38) which are both reduced by NADPH-dependent mechanisms and oxidized by H2O2-dependent mechanisms. The GSH/GSSG couple is maintained by a splice variant of GSSG reductase, which is targeted to mitochondria. Oxidation of GSH can occur by many enzymes, most notably glutathione peroxidase-1 (Gpx1) and -4 (Gpx4) and a number of GSH transferases (39). Glutaredoxin-2 also oxidizes GSH to GSSG in its reaction to remove GSH from glutathionylated proteins. 1. Remove treatment media from cells and wash three times using cold PBS. 2. Add approximately 500 mL of 1% digitonin (w/v) to each plate/ well of cells and incubate for 5 min on ice to permeablize.

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3. Aspirate the digitonin solution from the cells and carefully wash the cells three times with cold PBS. 4. Apply approximately 500 mL of 10% PCA/BA directly to the permeabilized, adherent cells, scrape each well and collect cell extracts in a 1.5-mL tube. 5. Samples are spun for 2 min in a microcentrifuge at approximately 11,000 ´ g to pellet protein. 6. An aliquot (300 ml) of each supernatant is transferred to a fresh microcentrifuge tube. 7. 9.3 mg/ml Iodoacetic acid solution (60 ml) is added to each tube and vortex to mix. 8. The pH is adjusted to 9.0 ± 0.2 with the KOH/tetraborate solution (approximately 220–250 ml). 9. After about 3 min to allow complete precipitation of potassium perchlorate, the pH of at least some of the samples should be checked to verify that they are in the correct range. 10. Incubate samples for 20 min at room temperature. 11. 300 ml of dansyl chloride (20 mg/ml acetone) is added, and the samples are mixed and placed in the dark at room temperature for 16–26 h. Dansylation of GSH is complete by 8 h, but GSSG has two amino groups that must be modified. The rate of the second dansylation is slower than that for the first with the result that mono-dansyl derivatives of GSSG (eluting between the N-dansyl-, S-carboxymethyl-GSH, and bisdansyl-GSSG) will be present if dansylation is incomplete. 12. After derivatization, chloroform (500 ml) is added to each tube to extract the unreacted dansyl chloride. 13. Samples are stored at 0–4° in the dark in the presence of both the perchlorate precipitate and the chloroform layer until assay by HPLC. Stability tests show that samples can be stored under these conditions for 12 months with little change in the amounts of GSH and GSSG derivatives. 3.1.2. HPLC Analysis of GSH/GSSG Redox

1. Samples are centrifuged for 2 min in a microcentrifuge prior to transfer of an aliquot of the upper (aqueous) layer to an autosampler. 2. Typical injection volume is 25 ml. 3. Separation is achieved on 3-aminopropyl columns (5 mm; 4.6 mm × 25 cm; Custom LC, Houston; or Sulpelcosil LC NH2, Supelco, Bellefonte, PA). 4. Initial solvent conditions are 80% A, 20% B run at 1 ml/min for 10 min. 5. A linear gradient to 20% A, 80% B is run over the period from 10 to 30 min.

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6. From 30 to 46 min, the conditions are maintained at 20% A, 80% B and returned to 80% A, 20% B from 46 to 48 min. 7. Equilibration time for the next run is 12 min. 8. Detection is obtained by fluorescence monitoring with bandpass filters, 305–395 nm excitation and 510–650 nm emission (Gilson Medical Electronics, Middleton, WI). 9. Quantification is obtained by integration relative to the internal standard, g-glutamylglutamate. Human Trx2 contains two Cys residues in the active site; the dithiol and disulfide forms are separated by nonreducing PAGE following modification of the protein with AMS, a reagent which increases the mass from 12 to 13 kDa (1 kDa/2 Cys) (Fig. 3). To obtain EhTrx2, a redox western blot method is used (40). This approach has an advantage over the GSH/GSSG method in that study of proteins which are present only in mitochondria provides a compartment-specific assay without subcellular fractionation. Redox western blots have been developed which resolve reduced and oxidized forms of proteins based upon both changes in molecular mass and changes is charge. For the former, a thiol reagent with relatively large mass, such as AMS (4-acetoamido-4¢-maleimidylstilbene-2,2¢-disulfonic acid), is used to increase the mass of the thiol-containing form of the protein. This allows separation by electrophoresis using SDS-PAGE, and is described below.

Trx2Red Trx2Ox E h Trx2 (mV)

3.2. Trx2 Redox Western Blot Analysis in Cultured Cells: Sample Preparation

-350 -300 -250 +Glc

None

Rot

AA

Stig

KCN

+Gln -Glc,-Gln Fig. 3. Trx2 redox state in HT29 cells exposed to mitochondrial respiratory inhibitors in control media and glucose- (Glc-), glutamine- (Gln-) free media. Cells were grown to 80% confluency and then cultured for 24 h with media as indicated. After 24 h, inhibitors were added at 5 mM (Rot, AA, Stig) or 0.5 mM (KCN) for 30 min prior to extraction and analysis by redox western blotting. Separation was provided by reaction of samples with AMS, a thiol reagent which adds approximately 500 Da per thiol, thereby slowing mobility sufficiently to separate the reduced form from the disulfide form. All incubations in Glc,-Gln-free media were significantly different from respective +Glc,+Gln controls. N = 5. Rot rotenone; AA antimycin A; Stig stigmatellin.

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3.2.1. Cell Lysis and Protein Alkylation

1. Assays are performed with cells grown in a 35-mm or 6-well culture plate (1–2 × 106 cells). 2. After experimental treatment, cells are washed with ice-cold PBS. 3. The cells are then treated with 1 mL of ice-cold TCA (10%), scraped, transferred to microcentrifuge tubes, incubated on ice for 30 min, and centrifuged at 12,000 × g for 10 min. 4. The supernatant is removed and the protein pellet is saved. 5. 1 mL of 100% acetone is added to the pellet. The tube is mixed, incubated on ice for 30 min, and centrifuged at 12,000 × g for 10 min. 6. The acetone is removed and the pellet (30–50 mg protein) is used for analysis. 7. After addition of 100 mL of lysis/derivatization buffer (50 mM Tris, pH 8.0, 0.1% SDS, 15 mM AMS), pellets are resuspended by sonication and incubated at room temperature in the dark for 3 h. The samples can be used for redox western blotting at this step or be saved at −20°C for subsequent analysis.

3.3. Prx3 Redox Western Blot Analysis in Cultured Cells: Sample Preparation

Peroxiredoxins (Prx) are a class of thiol-dependent antioxidant proteins that exhibit peroxidase activity. These proteins utilize redox-active Cys residues in their active site to reduce hydrogen peroxide and other organic peroxides. Prx are classified into two distinct groups, 1-Cys and 2-Cys, based on the number of Cys residues involved in the reduction of peroxides. Mammals possess six isoforms, five of which are considered 2-Cys Prx (Prx1-5). Prx 6 is the sole member of the 1-Cys class. The reaction mechanism for Prx occurs in two steps, the first of which is common for both 1-Cys and 2-Cys Prx. First, peroxide reacts with the “peroxidatic” Cys residue in the active site resulting in the formation of a cysteinesulfenic acid. This newly formed sulfenic acid then reacts with the “resolving” Cys, resulting in the formation of a disulfide linkage. It should be noted that Prx are present as domain-swapped homodimers, where the “peroxidatic” Cys is located on one subunit and “resolving” Cys on the opposite subunit. Therefore, when a 2-Cys Prx becomes oxidized the enzyme is locked in a dimer due to the newly formed disulfide bonds (Fig. 4). 2-Cys Prx utilize the thioredoxin/thioredoxin reductase system to reduce this disulfide and effectively recycle the enzyme back to its active form (41). Prx3 and Prx5 are mitochondrial isoforms; however, Prx5 is an “atypical” 2-Cys Prx that does not form an intermolecular disulfide upon reaction with a peroxide because it functions as a monomer. Based on the fact that Prx form stable disulfides resulting in a “locked” dimerized state, Cox et al. have exploited this concept in order to measure redox changes in Prx3 (42). This method involves alkylation of cellular proteins with NEM followed by SDS-PAGE and standard immunoblotting techniques.

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50kDa

265

37kDa

Prx3Ox (dimer)

25kDa

Prx3Red (monomer)

PQ, µM: Fig. 4. Measurement of oxidation of Prx3 by redox western blot analysis. SH-SY5Y human neuroblastoma cells were treated 24 h with increasing amounts of paraquat (0, 10, 25, 50 75, 100 mM). Following extraction and treatment with NEM to prevent further oxidation, separation by SDS-PAGE followed by western blotting reveals that the oxidized (disulfide) form increases in response to PQ compared to the reduced, NEM-modified form (monomer).

3.3.1. Cell Lysis and Protein Alkylation

1. Approximately 1–2 × 106 SH-SY5Y neuroblastoma cells are plated into each well of a six-well plate containing 2 mL of media and allowed to adhere overnight. 2. Media is aspirated from each well and replaced with treatment media containing desired concentrations of paraquat and cells are incubated for 24 h. 3. At the completion of the treatment period, the media is aspirated from each well and wells are gently washed three times with 2 mL of cold PBS. 4. After washing approximately 100 mL of alkylation buffer containing 100 mM NEM is added to each well and allowed to incubate 10–15 min at room temperature. 5. Add approximately 4 mL of 25% CHAPS solution to each well (1% final concentration) to lyse cells. 6. Using a cell scraper, scrape cells and debris from each well and collect cell extracts in 1.5 mL tubes and place on ice. 7. Using a benchtop sonicator, sonicate each tube briefly to ensure complete cell lysis and protein extraction. Keep cells on ice. 8. Pellet debris and insoluble protein by centrifugation at 14,000 rpm for 5 min on a benchtop microcentrifuge. Place samples back on ice. 9. Assay protein content of each sample using BCA protein assay (Thermo Scientific, Rockford, IL). 10. Add 5× nonreducing sample buffer to approximately 20–25 mg of protein. Place samples on a heating block at 95°C for 5 min.

3.4. TrxR2 Redox Western Blot Analysis: Sample Preparation

Trx system is composed of Trx, thioredoxin reductase (TrxR), NADPH, and Trx peroxidases/peroxiredoxins. Trx reduces protein disulfides directly and serves as a reductant for the peroxiredoxins (43). Oxidized form of Trx is reduced by catalytic activity of TrxR using an electron from reduced NADPH (44). TrxR2 is an isoform

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IP: Anti-TrxR2 WB: Streptavidin

+Glc

-Glc

-Gln

+Gln

-Gln

+AA

-TCEP +TCEP

WB : Anti-TrxR2

Fig. 5. Semiquantitative analysis of fractional reduction of thioredoxin reductase-2 (TrxR2) by BIAM-blot. Cells were incubated 24 h with +Glc,+Gln, −Glc,−Gln, or −Glc,−Gln and antimycin A (30 min with 5 mM; +AA). Aliquots of cell lysates were treated with the biotinylated iodoacetamide reagent, BIAM, and parallel aliquots were reduced with TCEP and then reacted with BIAM. Following immunoprecipitation with anti-TrxR2, samples were separated by SDS-PAGE, blotted and probed with fluorescently labeled streptavidin. Controls for recovery following immunoprecipitation were performed by western blotting with anti-TrxR2 and showed similar recovery. Although these methods are reproducible, the limiting conditions of BIAM labeling selected to maximize detection of reactive thiols in the presence of less reactive thiols does not allow strict quantification.

of TrxRs including E. coli TrxR, human TrxR1. The catalytic site of TrxR is -Cys-Val-Asn-Val-Gly-Cys- and located in the FAD domain of enzymes (45). Furthermore, TrxR2 has a C-terminal selenocysteine residue that is required for catalytic activity but is not part of the conserved active site (46). Since TrxRs are known to reduce oxidized Trx, alterations in TrxR activity may regulate Trx activity. TrxR2 is localized in mitochondria while TrxR1 is predominantly found in cytosol and nucleus (47). In this chapter, we describe an assay for measuring mitochondrial TrxR2 redox state using BIAM-labeling technique (Fig. 5). The thiol-reactive biotin iodoacetamide and biotin maleimide derivatives can be used for BIAM-labeling technique described in this chapter. This method is based on the procedure of Kim et al. (48), with modifications to measurement of redox states of proteins containing thiols in catalytic site such as TrxR and redox factor-1 (Ref-1). The procedures are described below. 3.4.1. Cell Culture and Affinity Purification

1. Cells (1–2 × 107 in 10 cm plate) after treatment [e.g., Glucose (Glc) and Glutamine (Gln) deficient media] were washed with cold PBS, fractionated, and then nuclear fraction was lysed with 1 ml of lysis buffer A [50 mM Bis–Tris–HCl (pH 6.5), 0.5% Triton X-100, 0.5% deoxycholate, 0.1% SDS, 150 mM NaCl, 1 mM EDTA, leupeptin, aprotinin, and 0.1 mM PMSF] containing 10 mM BIAM. As a control, cells that were not stimulated with treatment (e.g., Glc and Gln deficient media) were lysed and labeled with BIAM.

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2. After incubation for 10 min at 37°C in the dark, the labeling reaction was stopped by adding IAM to 5 mM. TrxR2 in the reaction mixtures was precipitated with the use of rabbit antibodies to TrxR2 and protein-G-sepharose (40 ml per sample). (Optional: Streptavidin-agarose can be used instead of TrxR2 antibody to immunoprecipitate all BIAM-labeled proteins). 3. Immunocomplex of BIAM-labeled TrxR2 (use of TrxR2 antibody) or other BIAM-labeled proteins (use of streptavidinagarose, 40 ml) are washed with 1 ml of ice-cold lysis buffer A three times. 4. Add 40 ml of 2× gel loading buffer to each sample, heat at 95°C for 10 min. 3.5. SDS-PAGE: Trx2, Prx3, TrxR2 Redox Western Analysis

1. These instructions assume the use of a BioRad Mini Protean II or III gel electrophoresis system. Clean each glass plate first with water, then 100% methanol prior to gel casting. 2. For Trx2 and Prx3: prepare a 1.5-mm thick, 15% resolving gel by mixing 2.3 mL water, 5 mL 30% acrylamide solution, 2.5 mL 1.5 M Tris–HCl (pH 8.8), 100 mL 10% SDS, 100 mL APS, and 4 mL TEMED in a 15-mL tube. Invert tube to mix, pour gel solution between glass plates, and allow gel to polymerize for approximately 45–60 min at room temperature. Note: Be sure to leave sufficient room for a stacking gel. Also, fill the remainder of the glass plate with water to ensure that the gel polymerizes with a clean, straight edge. 3. For TrxR2: Prepare a 1.5-mm thick, 10% resolving gel by mixing 4 mL water, 3.3 mL 30% acrylamide solution, 2.5 mL 1.5 M Tris–HCl (pH 8.8), 100 mL 10% SDS, 100 mL APS, and 4 mL TEMED in a 15-mL tube. Invert tube to mix, pour gel solution between glass plates, and allow gel to polymerize for approximately 45–60 min at room temperature. Note: Be sure to leave sufficient room for a stacking gel. Also, fill the remainder of the glass plate with water to ensure that the gel polymerizes with a clean, straight edge. 4. After polymerization of resolving gel, prepare and pour the 5% stacking gel by mixing the following in a 15-mL tube: 2.7 mL water, 0.67 mL 30% acrylamide solution, 0.5 mL 1.0 M Tris–HCl (pH 6.8), 40 mL 10% SDS, 40 mL APS, and 4 mL TEMED. Pour off the water layer on the resolving gel and add stacking gel to the top of the glass plates. Place in desired comb (10 or 15 well) and allow to polymerize for 15–30 min. 5. Assemble the gel apparatus. 6. Prepare 500 mL of running buffer by adding 50 mL 10× running buffer to 450 mL water. Then fill inner gel chamber and add remainder of buffer to outer chamber. Allow the assembly to

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sit for a few minutes to ensure that there are now leaks. If no leaks are discovered then samples may be loaded. 7. Load samples and the precision plus protein standards to individual wells. 8. Place lid on gel apparatus and connect to the Power Pac 200. Run the gel at 150 V for approximately 60–80 min or until the dye front runs off the gel. 3.6. Western Blotting: Trx2, Prx3 TrxR2 Redox Western Analysis

1. After proteins are separated on the SDS-PAGE gel, proteins need to be transferred to a nitrocellulose (or PVDF) membrane for immunoblotting. The procedure described here assumes the use of a BioRad semidry transfer apparatus. A “wet” transfer can also be conducted; however, this procedure is not described here. 2. Preincubate two filter papers and an appropriately cut nitrocellulose membrane in cold transfer buffer for approximately 10–15 min. 3. After SDS-PAGE, discard the running buffer and carefully pry apart the glass plates to free the gel. 4. Place a filter paper saturated with transfer buffer on the semidry transfer apparatus and use a serological pipette to roll out any trapped air bubbles. Next place the membrane on the filter paper and roll out any air bubbles. Place the gel on top of the membrane and orient it to your liking. Again, carefully roll out any air bubbles. Place the last filter paper on top the gel and roll out any air bubbles. Note: It is important to not introduce any air bubbles in the filter paper–membrane–gel–filter paper sandwich. These bubbles will prevent proper protein transfer and produce poor quality blots. 5. After the filter paper sandwich is made, assemble the transfer apparatus and connect it to the Power Pac 200. Run the transfer at 20 V for 60 min. Note: If you cast a thinner gel run the transfer for slightly shorter time period, i.e., 0.75 mm gels for 45 min at 20 V. 6. After the transfer is complete, place the membrane in 10–20 mL of blocking buffer and incubate on a shaker table for at least 30 min at room temperature. 7. Prepare primary antibody solution by adding 4 mL of peroxiredoxin 3 antibody to 10 mL of fresh blocking buffer (1:2,500 dilution); 10 mL of Trx2, or TrxR2 antibody to 10 mL of fresh blocking buffer (1:1,000 dilution). 8. Discard blocking solution and add the primary antibody solution to the membrane and incubate on a shaker table overnight at 4°C. 9. After primary antibody incubation, pour used antibody solution into a clean 15 mL tube and save for another use.

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10. Wash membrane three times for 15 min each in 10–20 mL of PBS-T. Discard wash buffer after every wash. 11. For Trx2 and TrxR2: Prepare secondary antibody solution immediately prior to use by adding 2 mL of goat antirabbit Alexa Fluor680 secondary antibody to 10 mL of blocking buffer (1:5,000 dilution). Protect this solution from light to avoid photobleaching of the fluorophore. 12. For Prx3: Prepare secondary antibody solution immediately prior to use by adding 2 mL of goat antimouse Alexa Fluor680 secondary antibody to 10 mL of blocking buffer (1:5,000 dilution). Protect this solution from light to avoid photobleaching of the fluorophore. 13. Add secondary antibody solution to the membrane and incubate 45 min in the dark at room temperature on a shaker table. 14. Wash membrane three times for 15 min each in 10–20 mL of PBS-T in the dark. Discard wash buffer after every wash. 15. After the final washes, carefully place the membrane between two Kim Wipes and allow the membrane to dry in the dark. 16. Once the membrane is dry, scan it on a Li-Cor Odyssey infrared scanner at 700 nm. Adjust the intensity, contrast, and brightness of the scan to optimize your result. 3.7. Mass Spectrometry-Based Redox Proteomics

3.7.1. Cell Lysis and Protein Collection

To measure the fractional reduction of mitochondrial proteins, we have used the Isotope-Coded Affinity Tag reagent (Applied Biosystems, CA) with a sequential treatment procedure designed to yield the ratio of reduced:oxidized forms of specific tryptic peptides, as measured by tandem mass spectrometry (Fig. 6) (49). Data from this approach show that many proteins have Cys residues which are partially oxidized under steady-state conditions. Consequently, application of the mass spectrometry-based redox proteomics method can be expected to considerably improve the understanding of redox signaling and control. Importantly, this approach allows detection of mitochondrial proteins even without mitochondrial isolation so that the oxidation of mitochondrial proteins can be determined within the context of other cellular compartments. The methods used for redox proteomic analyses of mitochondrial proteins are based upon those used for nuclear proteins (10). Analyses can be performed in three different ways, by studying isolated mitochondria, by using digitonin to permeabilize cells and remove contaminating cytoplasm, or by directly analyzing cells and selecting mitochondrial proteins from the larger list of cytoplasmic proteins and proteins from other subcellular compartments. 1. Cells [mouse aortic endothelial cells (MAEC), 1–2 × 106 in 3.5 cm plate] after treatment [e.g., reduced (−150 mV) or oxidized (0 mV) extracellular EhCySS] were washed with cold PBS.

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Fig. 6. Mass spectrometry-based analysis of fractional reduction of protein using ICAT reagents. Proteins are extracted and treated with the heavy ICAT reagent (H) to label thiols. Following removal of excess reagent, samples are treated with TCEP to reduce disulfides. The newly formed thiols are modified by treatment with the light reagent (L). Following tryptic digestion, analysis by LC-MS/MS allows calculation of fractional reduction from the H:L ratio.

2. Add 1 ml 10% TCA to cells, scrape, and collect lysates in a clean 1.5 ml tube. Precipitation of white protein pellets will be observed. 3. Place on ice for 30 min and spin for 10 min at 16,100 × g (4°C). 4. Remove the supernatant, add 1 ml acetone to protein pellet, voltex, and spin again at 16,100 × g for 10 min (4°C). 5. Remove the supernatant and air-dry pellet for 1–2 min. 6. Add 100 ml denaturation buffer (50 mM Tris, pH 8.5, 0.1% SDS) to pellet, resuspend pellet by sonication for 2 s on ice. 7. Perform protein assay. 8. Transfer 120 mg protein in denatruation buffer and add more denaturation buffer if needed to make 80 ml as total sample volume. 3.7.2. ICAT Labeling

1. Add 120 mg protein in 80 ml denatruation buffer prepared as above to 20 ml of heavy ICAT reagent (H). 2. Vortex and snap-centrifuge. 3. Incubate for 60 min at 37°C. 4. Add 10 ml 100% TCA to sample and voltex.

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5. Add additional 400 ml of 10% TCA to “4” and transfer all to a clean 1.5 ml tube. 6. Place on ice for 30 min. 7. Centrifuge at 16,100 × g for 10 min (4°C). 8. Remove the supernatant. 9. Wash protein pellet with 500 ml, 100% acetone. 10. Centrifuge at 16,100 × g for 10 min. 11. Remove the supernatant. 12. Add 80 ml denaturation buffer to pellet. 13. Add 2 ml TCEP to sample and sonicate for 2 s on ice. 14. Incubate for 20 min at 37°C. 15. Add 20 ml light ICAT reagent (L) reagent to samples. 16. Voltex and snap-centrifuge. 17. Incubate for 60 min at 37°C. 3.7.3. Trypsinization of ICAT-Labeled Proteins

1. Dissolve trypsin included in ICAT assay kit in 200 mL of MilliQ-H2O. 2. Add 200 mL of trypsin to a sample labeled with H- and L-ICAT reagents above. 3. Vortex and snap-centrifuge. 4. Incubate 12–16 h at 37°C. 5. Vortex and snap-centrifuge.

3.7.4. Cation Exchange

1. Place sample in 3 ml tube. 2. Add 2 ml of cation-exchange buffer (CEB)-Load to sample (one drop per second). 3. Vortex and snap-centrifuge. 4. Check pH; it should be around 2.5–3.3 – if not add more CEB-Load until pH is correct. 5. Assemble cartridge holder with cation exchange cartridge. 6. Equilibrate cartridge by injecting 2 ml CEB-Load (one drop per second). 7. Inject sample onto cartridge slowly (one drop per second). 8. Wash with 1 ml CEB-Load (one drop per second). 9. Elute peptides with 500 ml CEB-Elute (one drop per second). 10. Collect flow-through.

3.7.5. Cleaning and Storing Cation Exchange Cartridge

1. Inject 1 ml CEB to clean cartridge (one drop per second). 2. Inject 2 ml CEB before storing cartridge at 4°C.

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3.7.6. Purifying Peptides and Cleaving Biotin

1. Insert avidin cartridge into cartridge holder. 2. Equilibrate cartridge by injecting 2 mL of affinity buffer (AB)elute (one drop per second) discard waste. 3. Inject 2 mL AB-Load (one drop per second) discard waste. 4. Neutralize samples by adding 500 mL of AB, voltex, and snapcentrifuge. 5. Check pH; pH should be at 7 – if not add more AB until it is correct. 6. Vortex and snap-centrifuge. 7. Slowly inject sample onto cartridge (one drop per second). 8. Collect flow-through; This fraction contains unlabeled fragments and can be analyzed at a later time if necessary. 9. Inject an additional 500 mL of AB-Load and collect in the same tube (or a fresh tube) as step 8. 10. Inject 1 mL of AB (one drop per second)-first wash and discard flow-through. 11. Inject 1 mL of AB (one drop per second)-second wash and collect first 500 mL but discard the latter 500 mL. 12. Inject 1 mL of MilliQ-H2O and discard flow-through.

3.7.7. Eluting ICAT-Labeled Samples

1. Inject 800 mL AB-Elute (one drop per second). 2. Discard initial 50 mL. 3. Collect the remaining 750 mL in tube. 4. Vortex and snap-centrifuge.

3.7.8. Cleaning and Storing Avidin Cartridge

1. Inject 2 ml AB (one drop per second) to clean avidin cartridge. 2. Inject 2 ml AB and store at 4°C.

3.7.9. Cleaving Biotin

1. Evaporate samples in speed vacuum. 2. In another tube mix cleaving agents A (95 ml) and B (5 ml) in a ratio of 95:5. 3. Add 95 mL of cleaving reagent solution to each sample. 4. Vortex and snap-centrifuge. 5. Incubate for 2 h at 37°C. 6. Vortex and snap-centrifuge. 7. Evaporate in speed vacuum. 8. Send off for mass spectrometry analysis.

3.7.10. Mass Spectrometry Analysis of Redox ICAT

An Ultimate 3000 nanoHPLC system (Dionex) with a nanobore column (0.075 × 150 mm Pepmap C18 100 Ǻ, 3 mm, Dionex) is used with the LC eluent being directly sprayed into a QSTAR XL

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system using a nanospray source from Proxeon Biosystems. The data from each salt cut are combined and processed by ProteinPilot software (Applied Biosystems). All quantification is performed by the ProteinPilot V2.0.1 software using the Swiss-Prot database. Quantification for proteins of interest is manually validated by examination of the raw data. 3.7.11. Calculation of Protein Redox State

Table 1 shows an example of measuring protein reduction/oxidation state affected by changes in extracellular redox conditions (49). The ratios of reduced (H) to oxidized (L) thiols analyzed by mass spectrometry enables to calculate protein redox state e.g., % oxidation = [L/(H + L)] × 100. Thirty mitochondrial proteins were sorted out from the original data (49) to calculate redox states of mitochondrial proteins using this approach.

4. Notes Regarding Redox Analyses 1. To facilitate simultaneous measurement of CySS (typically >40 mM) and GSSG (typically <200 nM), two detectors with different sensitivity settings are used in series. Fluorometric detectors with monochromators set at 335 nm for excitation and 515 nm emission can also be used with equivalent results, but sensitivity is substantially less because the narrower bandwidths limit the intensity of both excitation and emission light. 2. To calculate the EhGSSG, use the Nernst equation as follows: Eh = Eo + RT / nF ln [GSSG] / [GSH]2 , where Eo for this equation is equal to −276 mV.

(

)

3. An alternative redox western approach is adapted from the original method of Holmgren and Fagerstedt (50) for E. coli Trx in which iodoacetate is used to add negative charges to thiols so that separation can be obtained under native gel conditions. 4. Trx2 resolves at approximately 10–15 kDa and care should be taken to prevent these lower mass proteins from running off of the gel. 5. Care must be taken to avoid loading too much protein because over-loading interferes with separation of reduced and oxidized Trx2. 6. The amount of protein that you load for each blot may be different depending on the cell type that is used. It is wise to optimize this condition to achieve the best result. 7. The mouse monoclonal antibody from Abcam is used in this current protocol; however, other antibodies against peroxiredoxin 3 exist and can be used if blotting conditions are optimized.

60 kDa heat shock protein, mitochondrial precursor

Acyl carrier protein, mitochondrial precursor

Adenylyl cyclase-associated protein 1

ADP/ATP translocase 1

ADP/ATP translocase 2

ADP-ribosylation factor 6

Aldehyde dehydrogenase, mitochondrial precursor

Aspartate aminotransferase, mitochondrial precursor

ATP synthase D chain, mitochondrial

ATP synthase subunit alpha, mitochondrial precursor

D-3-phosphoglycerate

Dehydrogenase/reductase SDR family member 8 precursor

Dihydrolipoyl dehydrogenase, mitochondrial precursor

P63038

Q9CR21

P40124

P48962

P51881

P62331

P47738

P05202

Q9DCX2

Q03265

Q61753

Q9EQ06

O08749

dehydrogenase

Name

Accession

2.0

2.9

2.9

3.8

4.1

1.5

1.8

2.2

4.0

2.7

2.2

3.1

4.9 2.9

1.2

2.3

3.5

1.3

2.9

1.1

0 mV H:L

1.5

2.5

4.7

2.3

3.9

1.9

−150 mV H:L

33.3

25.7

25.7

20.9

19.6

25.6

16.8

40.6

28.5

17.5

30.0

20.4

34.4

−150 mV % oxidation

39.7

35.1

31.5

20.1

26.7

30.9

24.4

45.5

30.5

22.3

42.9

25.7

46.9

0 mV % oxidation

Table 1 Redox ICAT results of mitochondrial proteins in mouse aortic endothelial cells treated with extracellular EhCySS of −150 mV (reduced) or 0 mV (oxidized)

6.4

9.4

5.8

0.8

7.1

5.3

7.6

4.8

2.0

4.8

12.9

5.3

12.6

D% oxidation

274 J.R. Roede et al.

Name

Dolichyl-diphosphooligosaccharideprotein glycosyltransferase 67 kDa subunit precursor

Electron transfer flavoprotein subunit alpha, mitochondrial precursor

Glutamate dehydrogenase 1, mitochondrial precursor

Isocitrate dehydrogenase [NADP], mitochondrial precursor

Malate dehydrogenase, mitochondrial precursor

Mitochondrial carrier homolog 2

NADH dehydrogenase [ubiquinone] iron-sulfur protein 6, mitochondrial precursor

NADH-cytochrome b5 reductase 3

Pyrroline-5-carboxylate reductase 2

Pyruvate kinase isozymes M1/M2

Succinate dehydrogenase [ubiquinone] flavoprotein subunit, mitochondrial precursor

Sulfide:quinone oxidoreductase, mitochondrial precursor

Accession

Q91YQ5

Q99LC5

P26443

P54071

P08249

Q791V5

P52503

Q9DCN2

Q922Q4

P52480

Q8K2B3

Q9R112

2.7

3.2

3.6

2.4

2.8

2.5

2.3

3.2

3.5

2.5

1.8

3.7

−150 mV H:L

1.9

2.3

2.0

2.0

2.1

1.9

3.1

2.3

1.2

1.8

1.7

0.8

0 mV H:L

27.0

23.8

21.6

29.4

26.5

28.7

30.1

23.8

22.4

28.5

35.2

21.3

−150 mV % oxidation

35.0

30.7

33.8

33.5

32.1

34.5

24.4

30.1

44.7

36.1

37.0

54.1

0 mV % oxidation

(continued)

8.1

6.9

12.2

4.1

5.5

5.8

5.7

6.3

22.2

7.6

1.9

32.8

D% oxidation

17 Redox Equivalents and Mitochondrial Bioenergetics 275

Name

Ubiquinol-cytochrome-c reductase complex core protein 1, mitochondrial precursor

Valyl-tRNA synthetase

Voltage-dependent anion-selective channel protein 1

Voltage-dependent anion-selective channel protein 2

Voltage-dependent anion-selective channel protein 3

Accession

Q9CZ13

Q9Z1Q9

Q60932

Q60930

Q60931

Table 1 (continued)

2.6

4.1

3.9

1.2

4.5

−150 mV H:L

1.1

1.2

2.8

0.9

3.3

0 mV H:L

27.6

19.7

20.5

45.2

18.3

−150 mV % oxidation

47.3

45.2

26.4

51.7

23.4

0 mV % oxidation

19.6

25.6

5.9

6.5

5.1

D% oxidation

276 J.R. Roede et al.

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8. It is very important to make sure that there are no air bubbles present in the filter paper-gel sandwich during the protein transfer step. These bubbles can obscure bands of interest in ruin an entire experiment. 9. If one chooses to utilize PVDF membrane instead of nitrocellulose, remember to activate the PVDF by wetting with 100% methanol prior to placing in transfer buffer. Also, DO NOT wet nitrocellulose with 100% methanol prior to use. 10. The semidry transfer technique has been chosen because it is far less cumbersome and easier to perform compared to the wet transfer technique. Wet transfer can be done if the investigator so wishes. 11. As a negative control for peroxiredoxin 3 oxidation, simply take an aliquot of your alkylated cell extract and add reducing sample buffer (containing DTT or b-mercaptoethanol). This will result in only one band (approximately 25 kDa) on the blot corresponding to a completely reduced protein.

5. Comments and Perspectives Electron transfer reactions between some of the components within the respiratory chain are very rapid, presenting a false impression that redox processes within the mitochondrion occur under near equilibrium conditions. In reality, the overall process is a nonequilibrium state, and partial rate control occurs at many steps. Similarly, proteins are synthesized with the thiol of Cys residues in the reduced state, presenting the impression that thiols are maintained in that form. However, the reality is that a fraction of the Cys residues are readily oxidized, and the steady-state generation of oxidants within mitochondria is sufficient to maintain many proteins in a partially oxidized state. An example is shown in Fig. 7 in which steady-state reduction of proteins in the pathway from NADH through TrxR2, Trx2, and Prx3 were examined under conditions which caused oxidation of NADPH. Results show that the entire pathway functions in a nonequilibrium steady-state. Of critical importance, the reactions which are not at equilibrium (within the electron transfer chain and among the protein thiols) are of most interest in terms of control of mitochondrial functions, and these are the most difficult to study because of the methodological challenges to accurately trap the steady-state values. Finally, an unresolved complexity lies in the relatively large number of modifications which thiols undergo under relevant biologic conditions. Irreversible modification occurs by reaction with the lipid oxidation product, 4-hydroxynonenal (51) and other reactive electrophiles. In addition, subsets of proteins are physiologically

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-Glc,-Gln

-Glc,-Gln,

+AA

100

Reduced Form (%)

80 60 40 20

Prx3

Trx2

TrxR2

NADPH

Prx3

Trx2

TrxR2

NADPH

Prx3

Trx2

TrxR2

NADPH

0

Fig. 7. Measurement of steady-state reduction of Trx2 by redox-western blotting and TrxR2 and Prx3 by BIAM blotting shows that mitochondrial thiol redox systems exist under nonequilibrium conditions in cells. The reduced fraction of TrxR2, Trx2, and Prx3 was decreased by both -Glc,-Gln media and by addition of respiratory substrates. Total cell NADPH was measured by HPLC and is shown for comparison. Results are representative of four experiments.

regulated by glutathionylation and nitrosylation (52). The assays described above do not discriminate the contributions of these other modifications. Consequently, accurate descriptions of mitochondrial function will require more global approaches which capture the contributions of each rate-controlling step.

Acknowledgements Supported by NIH grants ES009047, ES011195, and ES012870. References 1. Mitchell P (1979) Keilin’s respiratory chain concept and its chemiosmotic consequences. Science 206:1148–1159 2. Chance B, Sies H, Boveris A (1979) Hydroperoxide metabolism in mammalian organs. Physiol Rev 59:527–605 3. Meredith MJ, Reed DJ (1982) Status of the mitochondrial pool of glutathione in the isolated hepatocyte. J Biol Chem 257:3747–3753

4. Wallace DC (1999) Mitochondrial diseases in man and mouse. Science 283:1482–1488 5. Jones DP (2006) Disruption of mitochondrial redox circuitry in oxidative stress. Chem Biol Interact 163:38–53 6. Chance B (1957) Cellular oxygen requirements. Fed Proc 16:671–680 7. Taylor ER, Hurrell F, Shannon RJ, Lin TK, Hirst J, Murphy MP (2003) Reversible

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glutathionylation of complex I increases mitochondrial superoxide formation. J Biol Chem 278:19603–19610 Zhang R, Al-Lamki R, Bai L et al (2004) Thioredoxin-2 inhibits mitochondria-located ASK1-mediated apoptosis in a JNK-independent manner. Circ Res 94:1483–1491 Lillig CH, Berndt C, Vergnolle O et al (2005) Characterization of human glutaredoxin 2 as iron-sulfur protein: a possible role as redox sensor. Proc Natl Acad Sci USA 102:8168–8173 Go YM, Pohl J, Jones DP (2009) Quantification of redox conditions in the nucleus. Methods Mol Biol 464:303–317 Jones DP (1984) Effect of mitochondrial clustering on O2 supply in hepatocytes. Am J Physiol 247:C83–C89 Schafer FQ, Buettner GR (2001) Redox environment of the cell as viewed through the redox state of the glutathione disulfide/glutathione couple. Free Radic Biol Med 30:1191–1212 Chance B (1954) Spectrophotometry of intracellular respiratory pigments. Science 120:767–775 Keilin D (1966) The history of cell respiration and cytochrome. Cambridge University Press, Cambridge Chance B (1952) Spectra and reaction kinetics of respiratory pigments of homogenized and intact cells. Nature 169:215–221 Jones DP, Thor H, Andersson B, Orrenius S (1978) Detoxification reactions in isolated hepatocytes. Role of glutathione peroxidase, catalase, and formaldehyde dehydrogenase in reactions relating to N-demethylation by the cytochrome P-450 system. J Biol Chem 253:6031–6037 Tamura M, Hazeki O, Nioka S, Chance B (1989) In vivo study of tissue oxygen metabolism using optical and nuclear magnetic resonance spectroscopies. Annu Rev Physiol 51:813–834 Jones DP (1981) Determination of pyridine dinucleotides in cell extracts by high-performance liquid chromatography. J Chromatogr 225: 446–449 Williamson JR, Corkey BE (1969) Assays of intermediates of the citric acid cycle and related components by fluorometric enzyme methods. Methods Enzymol 13:434–513 Sies H (1982) Nicotinamide nucleotide compartmentation. In: Sies H (ed) Metabolic compartmentation. Academic, London, pp 205–231 Kirwan GM, Coffey VG, Niere JO, Hawley JA, Adams MJ (2009) Spectroscopic correlation analysis of NMR-based metabonomics in exercise science. Anal Chim Acta 652:173–179 Beylot M, Beaufrère B, Normand S, Riou JP, Cohen R, Momex R (1986) Determination of human ketone body kinetics using stableisotope labelled tracers. Diabetologia 29:90–96

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23. Jones DP, Kennedy FG (1982) Intracellular oxygen supply during hypoxia. Am J Physiol 243:C247–C253 24. Chance B, Schoener B (1962) Correlation of oxidation-reduction changes of intracellular reduced pyridine nucleotide and changes in electroencephalogram of the rat in anoxia. Nature 195:956–958 25. Chance B, Cohen P, Jobsis F, Schoener B (1962) Intracellular oxidation-reduction states in vivo. Science 137:499–508 26. Song Y, Buettner GR (2010) Thermodynamic and kinetic considerations for the reaction of semiquinone radicals to form superoxide and hydrogen peroxide. Free Radic Biol Med 49(6):919–962 27. Yamamoto Y, Yamashita S (2002) Ubiquinol/ ubiquinone ratio as a marker of oxidative stress. Methods Mol Biol 186:241–246 28. Matsubara M, Ranji M, Leshnower BG et al (2010) In vivo fluorometric assessment of cyclosporine on mitochondrial function during myocardial ischemia and reperfusion. Ann Thorac Surg 89:1532–1537 29. Scholz R, Thurman RG, Williamson JR, Chance B, Bucher T (1969) Flavin and pyridine nucleotide oxidation-reduction changes in perfused rat liver. I. Anoxia and subcellular localization of fluorescent flavoproteins. J Biol Chem 244:2317–2324 30. Tamura M, Oshino N, Chance B, Silver IA (1978) Optical measurements of intracellular oxygen concentration of rat heart in vitro. Arch Biochem Biophys 191:8–22 31. Estabrook RW (1961) Studies of oxidative phosphorylation with potassium ferricyanide as electron acceptor. J Biol Chem 236: 3051–3057 32. Jones DP, Orrenius S, Mason HS (1979) Hemoprotein quantitation in isolated hepatocytes. Biochim Biophys Acta 576:17–29 33. Aw TY, Andersson BS, Jones DP (1987) Suppression of mitochondrial respiratory function after short-term anoxia. Am J Physiol 252:C362–C368 34. Jones DP (1982) Intracellular catalase function: analysis of the catalytic activity by product formation in isolated liver cells. Arch Biochem Biophys 214:806–814 35. Guidot DM, Repine JE, Kitlowski AD et al (1995) Mitochondrial respiration scavenges extramitochondrial superoxide anion via a nonenzymatic mechanism. J Clin Invest 96: 1131–1136 36. Jones DP (2006) Redefining oxidative stress. Antioxid Redox Signal 8:1865–1879 37. Sies H, Jones DP (2007) Oxidative stress. In: Fink G (ed) Encyclopedia of stress, 2nd edn. Elsevier, New York, pp 45–48

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38. Zhang H, Go YM, Jones DP (2007) Mitochondrial thioredoxin-2/peroxiredoxin-3 system functions in parallel with mitochondrial GSH system in protection against oxidative stress. Arch Biochem Biophys 465:119–126 39. Jones DP (2008) Radical-free biology of oxidative stress. Am J Physiol Cell Physiol 295:C849–C868 40. Halvey PJ, Watson WH, Hansen JM, Go YM, Samali A, Jones DP (2005) Compartmental oxidation of thiol-disulphide redox couples during epidermal growth factor signalling. Biochem J 386:215–219 41. Wood ZA, Schroder E, Robin Harris J, Poole LB (2003) Structure, mechanism and regulation of peroxiredoxins. Trends Biochem Sci 28:32–40 42. Cox AG, Winterbourn CC, Hampton MB (2010) Measuring the redox state of cellular peroxiredoxins by immunoblotting. Methods Enzymol 474:51–66 43. Padgett CM, Whorton AR (1995) S-nitrosoglutathione reversibly inhibits GAPDH by S-nitrosylation. Am J Physiol 269:C739–C749 44. Stadtman TC (2002) Discoveries of vitamin B12 and selenium enzymes. Annu Rev Biochem 71:1–16

45. Gasdaska PY, Gasdaska JR, Cochran S, Powis G (1995) Cloning and sequencing of a human thioredoxin reductase. FEBS Lett 373:5–9 46. Mustacich D, Powis G (2000) Thioredoxin reductase. Biochem J 346(Pt 1):1–8 47. Soini Y, Kahlos K, Napankangas U et al (2001) Widespread expression of thioredoxin and thioredoxin reductase in non-small cell lung carcinoma. Clin Cancer Res 7:1750–1757 48. Kim JR, Lee SM, Cho SH et al (2004) Oxidation of thioredoxin reductase in HeLa cells stimulated with tumor necrosis factoralpha. FEBS Lett 567:189–196 49. Go YM, Park H, Koval M et al (2010) A key role for mitochondria in endothelial signaling by plasma cysteine/cystine redox potential. Free Radic Biol Med 48:275–283 50. Holmgren A, Fagerstedt M (1982) The in vivo distribution of oxidized and reduced thioredoxin in Escherichia coli. J Biol Chem 257:6926–6930 51. Roede JR, Jones DP (2010) Reactive species and mitochondrial dysfunction: mechanistic significance of 4-hydroxynonenal. Environ Mol Mutagen 51:380–390 52. Requejo R, Chouchani ET, Hurd TR, Menger KE, Hampton MB, Murphy MP (2010) Measuring mitochondrial protein thiol redox state. Methods Enzymol 474:123–147

Chapter 18 NMR Methodologies for Studying Mitochondrial Bioenergetics Tiago C. Alves, Ivana Jarak, and Rui A. Carvalho Abstract Nuclear magnetic resonance (NMR) spectroscopy is a technique with an increasing importance in the study of metabolic diseases. Its initial important role in the determination of chemical structures (1, 2) has been considerably overcome by its potential for the in vivo study of metabolism (3–5). The main characteristic that makes this technique so attractive is its noninvasiveness. Only nuclei capable of transitioning between energy states, in the presence of an intense and constant magnetic field, are studied. This includes abundant nuclei such as proton (1H) and phosphorous (31P), as well as stable isotopes such as deuterium (2H) and carbon 13 (13C). This allows a wide range of applications that vary from the determination of water distribution in tissues (as obtained in a magnetic resonance imaging scan) to the calculation of metabolic fluxes under ex vivo and in vivo conditions without the need to use radioactive tracers or tissue biopsies (as in a magnetic resonance spectroscopy (MRS) scan). In this chapter, some technical aspects of the methodology of an NMR/MRS experiment as well as how it can be used to study mitochondrial bioenergetics are overviewed. Advantages and disadvantages of in vivo MRS versus high-resolution NMR using proton high rotation magic angle spinning (HRMAS) of tissue biopsies and tissue extracts are also discussed. Key words: NMR spectroscopy, Krebs cycle, Intermediary metabolism, Isotopomer analyses, Stable isotopes

1. Introduction: Mitochondrial Metabolism

The word metabolism has its origin in the Greek “metabole” which means “change.” Therefore, in a physiological context, metabolism is a term used to characterize the set of chemical reactions able to convert specific molecules into different ones for either energy production or synthesis of biomolecules required for structural and replication purposes. In particular, the production of energy, supported by the oxidative metabolism, assumes special importance since it is vital for cell sustainability.

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_18, © Springer Science+Business Media, LLC 2012

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Mitochondria play an important role in this since it is the organelle in which 95% of the production of energy occurs. Two main substrates, glucose and fatty acids, are used to extract energy from (Eqs. 1 and 2, respectively). Once in the cells, glucose is metabolized into pyruvate through a series of enzymatic catalyzed reactions occurring in the cytosol. With this glycolytic process only two molecules of ATP and two of NADH are produced per molecule of glucose. Therefore, in order to extract more energy, pyruvate needs to be further oxidized. In the matrix of mitochondria, pyruvate goes through a process of oxidative decarboxylation, mediated by pyruvate dehydrogenase (PDH), originating acetylCoA and NADH. + + Glucose + 2NAD + 2Pi + 2ADP ® 2Pyruvate + 2NADH + 2ATP + 2H + 2H 2O

(1)

+ + Palmitate + 7NAD + 7FAD ® 8Acetyl - CoA + 7NADH + 7FADH 2

(2)

Also fatty acids, after the activation and transport into the mitochondria, are oxidized through a series of reactions, known as b-oxidation, that culminate in the production of acetyl-CoA, NADH, and FADH2. These are key metabolites in intermediate metabolism because they serve as links between the oxidation of substrates and the production of energy. This happens by two main processes, oxidation of acetyl-CoA through the Krebs cycle (TCA) and ATP synthesis coupled to the oxidation of NADH and FADH2 (Fig. 1). During the first process acetyl-CoA is condensed with oxaloacetate to produce citrate which, in turn, undergoes further oxidation regenerating oxaloacetate and producing, as a result, NADH and FADH2. These, in turn, are the reducing equivalents that allow the production of ATP. The inner membrane of the mitochondria contains a series of protein complexes – electron transport chain – involved in the acceptance and transport of electrons from these species. Two electrons are transferred from NADH to the NADH dehydrogenase complex (also known as complex I) or from FADH2 to the succinate dehydrogenase complex (complex II). The electrons are then transferred successively to complexes III and IV. Oxygen is the final acceptor of these electrons being converted to water. The passage of the electrons through complexes I, III, and IV leads to transport of H+ from the matrix to the intermembrane space creating an electrochemical gradient. This potential energy is then used to synthesize ATP. The transport of these H+ through the ATP synthase complex (complex V) back to the mitochondrial matrix provides the necessary mechanism to synthesize ATP from ADP and inorganic phosphate (Pi) (4). Due to its central role in the production of energy, dysfunctional mitochondrial metabolism leads to several metabolic disorders. A very common condition is the development of Type 2 Diabetes

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283

Cytasol H+ H+

ePyruvate

+ CoA+NAD

NADH

Ι

H+

ΙΙ

III

FADH2

co2 +NADH Acetyl-CoA

e-

Fatty Acid

Intermembrane Space

IV

Mitochondrial Matrix

H+ e-

H+

H+ O2

Citrate

H+

H2O ADP+Pi V

H+

Isocitrate

Oxaloacetate

ATP NAD+

NADH NAD+

NADH+CO2

Tricarboxylic Acid Cycle

Malate

a-Ketoglutrate NAD++CoA NADH+CO2 Succinyl-CoA

Fumarate

GDP +Pi

FADH2 FAD2+

Succinate

GTP+CoA

Fig. 1. Oxidative metabolism coupled to the synthesis of ATP in mitochondria. In tissues, glucose and fatty acids are metabolized to acetyl-CoA which in turn is further oxidized in the Krebs cycle (TCA) producing NADH and FADH2. These serve as electron donors to complexes I and II, respectively, and their passage through the electron transport chain provides the transport of protons (H+) from the matrix to the intermembrane space creating the electrochemical gradient necessary for ATP synthesis.

(T2D). Several studies performed in insulin-resistant individuals revealed several problems related with the mitochondrial genetic material, lower content of the electron transport chain complexes, and even reduced mitochondrial density (5–7). The consequences of these changes in metabolism are profound and can be followed using the MRS technique. In an elegant study, Petersen et al. (8) using 31P MRS determined that skeletal muscle of insulin-resistant offspring of T2D was characterized by a reduced rate of ATP synthesis. Also, using 1H MRS, this was associated with increased intramyocellular concentrations of triglycerides suggesting that the impaired mitochondrial metabolism was associated with a reduced oxidative capacity leading to accumulation of fatty acids. Several approaches can be undertaken for monitoring mitochondrial metabolism using nuclear magnetic resonance (NMR)based methodologies. The most obvious is certainly the evaluation of the production of ATP and other 31P-containing metabolites by in vivo 31P MRS and/or 31P NMR of tissue extracts. As mentioned

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above, one of the major functions of mitochondria is the conversion of reducing equivalents into ATP using the electron transport chain and the ATP synthase. Thus measurements of the rates of ATP synthesis and conversion of ATP into metabolites such as PCre are crucial for an adequate evaluation of mitochondrial metabolism. The performance of the TCA cycle can also be directly evaluated by 13C isotopomer analysis either in vivo by 13C MRS or by 13C NMR of tissue extracts. A kinetic evaluation of 13C incorporation in Krebs cycle intermediates or metabolites in fast exchange with them can be made by the administration of 13C-enriched oxidative substrates (9–11). Mitochondrial metabolism can also result in the alterations of the concentrations of several metabolic intermediates. An intricate interplay characterizes the equilibrium between cytosolic and mitochondrial metabolite pools, and 1H MRS as well as 1H NMR of tissue extracts can be used to follow the alterations in the intermediate metabolite pools. In some circumstances, an integration of the information derived from each of the above-mentioned approaches is crucial for an adequate evaluation of the metabolic status of a tissue or organ and infer about mitochondrial abnormalities. 1.1. In Vivo Magnetic Resonance Spectroscopy Experimental Setup

The ability to measure in vivo fluxes through oxidative and biosynthetic pathways is crucial to characterize the metabolic phenotype of a particular tissue in a certain disorder context. Due to its noninvasive character, in vivo magnetic resonance spectroscopy (MRS) assumes a key role in this task since metabolic rates can be quantified through the detection of naturally occurring nuclei such as 1H, 13 C, and 31P. To understand how an in vivo MRS experiment is conducted, each of the components involved in this process is briefly overviewed in the following sections.

1.1.1. Magnets

Every nucleus with a magnetic momentum has the ability to precess about an external magnetic field with a certain energy associated. As in all types of spectroscopy, the signal obtained is dependent on the transition between different levels of energy. In an MRS experiment, the difference between energy levels is proportional to the intensity of the magnetic field (Fig. 2a), and therefore, the higher the intensity of the external magnetic field the higher the sensitivity for the detection of any given NMR active nucleus. For this reason, the magnets used in this technique are built to produce magnetic fields several times more intense than the one produced by Earth. The typical field strengths used in human studies vary from 3 to 7 Tesla (T) while for animal studies they can go safely up to 9.4 T (Fig. 3) and even 14.1 T, which are several fold increases comparing to earth’s magnetic field (5 × 10−5 T). The modern magnets are able to achieve high-magnetic field intensities due to the development of superconducting technology which allows the circulation of a very high-electrical current density with a small resistance. The use of superconducting materials allows

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Fig. 2. (a) Variation of the energy difference (DE ) between the two energy levels that nuclear spins can assume with the intensity of the external magnetic field (B0). (b) Representation of the nuclear spin orientation in the presence of a B0. Precession along B0 corresponds to the lowest energy state (a) while the precession against B0 is the state of highest energy (b).

Fig. 3. Horizontal 9.4 T magnet used in animal studies.

also the generation of linear and stable magnetic fields that persist for a very long time. To achieve the superconducting properties of the magnet, these materials are kept at extremely low temperatures (below the boiling temperature of helium, ~4 K). The materials used are wind multifilamentary of niobium–titanium conductors embedded in a copper matrix for field strengths of 9 T. For higher fields, niobium–tin is also used.

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The magnets used for the in vivo studies are built to have a bore in the center large enough to accommodate the entire animal/ person. A moving platform is also common to carry the region of interest to the center of the magnet where the linearity and homogeneity of the magnetic field are the highest. An important consideration has to do with the appropriate location of the magnet due to magnetic fields outside the magnet. An unshielded 1.5 T magnet can extend the magnetic field until 10 m in all directions. The shielding is obtained through the use of extra coils on the outside of the magnet that counteract and reduce the external magnetic field. However, this procedure severely affects the linearity and homogeneity of the magnetic field in horizontal magnets. Therefore, the solution has been to place them in a compartment with steel walls which is very effective in limiting the magnetic field. 1.1.2. Radiofrequency Emitting/Receiving Probes

As shown in Fig. 2a, when in the presence of a strong B0, there are two energy levels among which the nuclear spins are distributed. In this situation, the nuclear spins in both states will precess about B0, either with the same or opposite direction of the magnetic field, with a defined frequency – Larmor frequency (Eq. 3). wo = gB0

(3)

This precession frequency is associated with energy and when the sample is irradiated with energy that equals the difference between the two levels, the nuclear spins transition from a to b state. The energy difference between the two states is smaller than in other types of spectroscopy and therefore irradiation with energy in the radio frequency (RF) range is able to promote the transition between states. This is accomplished by the use of a current oscillator. Simply, a coil of wire is placed along the x-axis (Fig. 4a); the passage of electrical current through this coil produces an oscillating magnetic field along the x-axis (B1), perpendicular to B0 (Fig. 4b). The effect of these perpendicular magnetic fields on the nuclear spins is crucial to the MRS experiments. In the presence of B0, the distribution of spins occurs according to Eq. 4, N b / N a = e -wo/ kT

(4)

in which N is the number of spins in each state, k is Boltzmann’s constant, 1.3805 × 10−23 J/K and T is the temperature in Kelvin. According to Eq. 4, at room temperature, the number of spins in the fundamental state is higher than in the upper energy state. Therefore, the sum of the magnetization vectors results in a positive net vector with the same direction as B0 (Mz). In this situation, because every spin have different precession frequencies, the component of the net magnetization vector on the xy plane (Mxy) cancels (Fig. 4a). In the presence of B1, all the spins precess at the

18

a

NMR Methodologies for Studying Mitochondrial Bioenergetics

b

Z

c

Z

287

y Receiving Coil

Mz Mxy

NMR Coil

y

B1

My

y X

X

X

Fig. 4. Mechanism of nuclear spin excitation by a RF pulse B1. (a) In the presence of an external magnetic field the sum of the magnetic vectors from both states results in a net magnetization vector with only a component along the z axis (Mz ). (b) In the presence of B1, the synchronization of precession frequencies and the transition of spins from a to b state, leads to a reduction of the Mz component and the emergence of Mxy. (c) Effect of the absence of B1 on the intensity of Mxy with time.

same frequency. In this situation, the alignment of nuclear spins promotes the emergence of the Mxy component of net magnetization vector oscillating in xy plane as long as B1 is on. At the same time, the component Mz is reduced as the spins from the fundamental state are excited to the upper state (Fig. 4b). As soon as the RF pulse is turned off, the nuclear spins start to dephase according to their chemical environment. As consequence of this dephasing the component Mxy of the net magnetization vector decreases with time (Fig. 4c). The alternate electromagnetic component on the xy plane is then detected by a receiving coil inducing an electric current that is detected and manipulated to acquire the signal. There are two main types of coils used in vivo. A volume coil, characterized by a coiled wire that completely surrounds the region of interest, and surface coils localized on the surface of the region of interest. Even though the volume coil provides a high homogeneity of the B1 field across the sample the sensitivity is low. In contrast, the surface coils provide high sensitivity; however, with this type of coil, the B1 field is more inhomogeneous leading to signal loss across. This can be overcome using localized spectroscopy in which spectra will be acquired from a voxel set in the most homogeneous region, typically on the surface of the region of interest. In Fig. 5 an example of a set up used in an in vivo experiment is presented. In this set up, used to measure in vivo ATP synthesis in skeletal muscle, it is possible to observe that the animal is anesthetized in a platform in a way that allows evidencing the region of interest while the physiological parameters are being monitored (Fig. 5a). When this is ready the platform is placed in a probe (Fig. 5b) in which the coils necessary for the experiment are localized (Fig. 5c).

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a

ECG Wire Anesthesia Carrier

Face Mask

Region of Interest

b

Animal Platfrom

Surface Coils

31

Coil

1H

Coil

Fig. 5. Example of an experimental set up for an in vivo MRS experiment: (a) set up where the animal is placed in a platform under anesthesia and physiological monitoring; (b) sliding of the platform in the probe in which the region of interest in placed under; (c) the coils (1H and 31P) used in the experiment.

1.2. 1H MRS

Proton MRS (1H MRS), due to the ubiquitous distribution of the 1H nuclei in intermediate metabolites and its intrinsic high NMR sensitivity, constitutes a very robust technique for the identification and quantification of metabolites in vivo. Several aspects, however, pose significant problems in its application. These include essentially a need for very efficient water suppression, since tissue water concentrations are several orders of magnitude higher than any of the other metabolites, significant resonance overlap due to the low chemical shift range, and also the impossibility of measuring low-concentration metabolites, frequently the most relevant in several metabolic pathways. In relation to mitochondrial oxidative metabolism, under most circumstances levels of Krebs cycle intermediates (e.g., a-ketoglutarate, oxaloacetate) are too small to allow their direct observation by 1H MRS and the evaluation of Krebs cycle fluxes is essentially made by observation of metabolic intermediates in exchange with the Krebs cycle intermediates, namely glutamate. Figure 6 shows an in vivo 1H MRS spectra from the hippocampus of a rat with Type I diabetes (steptozotocin induced). Several metabolites could be identified including lactate (Lac), alanine (Ala), g-amino-butyric acid (GABA), N-acetyl-aspartate (NAA), N-acetyl-aspartyl-glutamate (NAAG), glutamate (Glu),

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Fig. 6. Representative in vivo 1H NMR spectra expanded from 0.5 to 5.5 ppm obtained in the hippocampus of a 12-week-old STZ-induced diabetic rat. The spectra was measured by the SPECIAL (35) sequence with echo time of 2.8 ms, repetition time of 4 s, 640 scans, and VOI of 18 µL located in the hippocampus. For resolution enhancement, a shifted Gaussian function (gf = 0.12 and gsf = 0.05) was applied before Fourier transformation. Zero-phase but not baseline was corrected.

glutamine (Gln), aspartate (Asp), creatine (Cre), phosphocreatine (PCre), phosphocholine (PCho), glycerophosphocholine (GPC), taurine (Tau), inositol (Ins), and glucose (Glc). Changes in metabolite levels by themselves might be of little importance in studies of metabolic pathway dynamics. Thus, it is frequent to associate the high sensitivity of 1H MRS detection with the dynamic metabolic analysis afforded by the administration of 13C-enriched substrates. The administration of 13C enriched substrates leads to the enrichment of the intermediates of the metabolic pathways metabolizing them and could be monitored by conventional 1H MRS providing that the 13C satellites are well resolved (e.g., lactate, glucose) or by 1H-observed 13C-edited (POCE) NMR spectroscopy (12, 13). This allows the kinetic evaluation of 13C incorporation in metabolic intermediates and the modeling of metabolic pathways using suitable mathematical models (14, 15). More well-resolved 1H NMR spectra are always possible to acquire using 1H HRMAS of tissue biopsies (Fig. 7), or high-resolution 1H NMR of tissue extracts (e.g., perchloric acid extracts, Fig. 8). However, these spectra are much less noteworthy due to the invasive nature of both approaches. 1.3. 31P MRS

With a natural abundance of 100% and a high sensitivity (around 7% of the intensity of 1H), 31P assumes a very important role in the use of MRS to study metabolism. Also relevant in MRS is the resolution of the resonances of the metabolites of interest. A higher chemical shift dispersion allows a better identification and quantification of the resonances present in the spectrum. A typical 31P

Fig. 7. 14.1 T 1H HRMAS spectrum of a rat hippocampus. Resonances due to Lipids (L1, L2, L3), g-amino-butyric acid (GABA), N-acetyl-aspartate (NAA), N-acetyl-aspartyl-glutamate (NAAG), glutamate (Glu), glutamine (Gln), aspartate (Asp), creatine (Cre), phosphocreatine (PCre), phosphocholine (PCho), glycerophosphocholine (GPC), taurine (Tau), and inositol (Ins) are well resolved.

12+13CH -Lac 3 12+13C-H1α-GIc

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Fig. 8. 14.1 T 1H-NMR spectrum from the perchloric acid (PCA) extract of hippocampus slices superfused with [U-13C]glucose and [2-13C]acetate with (top) and without (bottom) 13C broadband decoupling. Expansions show the effect of 13C broadband decoupling on glucose H1a and lactate and alanine-CH3 resonances.

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Fig. 9. Typical 31P spectrum obtained from skeletal muscle extracts analyzed in a 11.7-T vertical bore magnet. The main resonances correspond to inorganic phosphate (Pi), phosphocreatine (PCr), and the ATP phosphate nuclei in the positions a, b, and g.

spectrum (Fig. 9) shows that all the resonances are dispersed within 30 ppm of spectral width. Another particularity of 31P NMR spectroscopy is the high sensitivity to factors like pH and complexation with ions like magnesium. At pH 6.5 and full magnesium complexation, Pi resonates at 5.7 ppm, g-ATP at −2.5 ppm, a-ATP at −7.7 ppm, and b-ATP at −18 ppm. Phosphocreatine (PCre), for being one of the most insensitive metabolites to both pH and magnesium concentrations, is frequently used as a reference (0 ppm). Other 31P metabolites are also identifiable in highly resolved 31P NMR spectra of tissue extracts (e.g., NADH, NADPH, ADP) but the same is frequently not true for in vivo 31P MRS spectra due to the field inhomogeneities that characterize tissues. Another possible use of 31P NMR spectroscopy is in the measurement of reaction kinetics involving phosphorus containing metabolites. Since the first report by Brindle et al. (16) of the use of 31P saturation transfer (Fig. 10) in skeletal muscle, this has emerged as a powerful technique to measure the rate of ATP synthesis in vivo (17–19). It is based on the assumption that a perturbation of the magnetization of one metabolite can be transferred to other through chemical exchange without affecting the rate of this reaction. The main route to synthesize ATP is through the phosphorylation of ADP in the mitochondria. This reaction can be described as in Eq. 5 where k1 and k−1 are the unidirectional rate constants for reactions of degradation and synthesis of ATP, respectively.

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? − ATP Saturation Symmetrical Saturation

? MPI

Fig. 10. Representative spectra obtained from a saturation transfer experiment. A first spectrum is acquired in which g-ATP is saturated. A control experiment is also performed in which the saturating frequency is set down-field equidistant from Pi as the g-ATP. DMPi is the difference of the Pi intensity between the two experiments.

k1

ATP  ADP + Pi k-1

(5)

Based on Eq. 5, the rate of ATP synthesis (VATP) is given by Eq. 6 where [·] represents the intracellular concentration of ADP and Pi. Because at steady state the concentration of ADP can be assumed to be constant so that k¢−1 = k−1[ADP]. V ATP = k-1 [ADP][Pi] » k-¢1 [Pi]

(6)

To use 31P MRS to determine VATP, the equilibrium shown in Eq. 5 must be described in terms of changes in the magnetization (i.e., intensity of the MR signal). Therefore, using the Bloch equations to express the variation of the longitudinal magnetization (Mz) with time, we have dM Pi = (M 0 Pi - M Pi (t )) - k-¢1M Pi (t ) - k1M ATP (t ) dt ® T1

(7)

Pi

in which MPi(t) and MATP(t) represent the longitudinal magnetization at a certain time point while MPi0 is the longitudinal magnetization at

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equilibrium; T1Pi is the longitudinal relaxation time constant for Pi. During a saturation transfer experiment, the resonance of g-ATP is saturated. Mechanistically, this is achieved through the use of a very long (~10 s) low power pulse that scrambles all spins of the targeted nucleus resulting in the loss of signal. The long pulse used allows also the chemical exchange to occur so that the saturated phosphate in the position g of the ATP is transferred to the pool of Pi (Eq. 5). At this point, because the Pi pool contains both “MR-visible” and “MR-invisible” Pi the total signal obtained is reduced when compared to a scan with no saturation. An important factor to take into account is that the saturation of a particular frequency is never perfect and its effects can be extended to other frequencies such as Pi. For this reason, a second experiment is performed in which the saturation is performed at a down-field frequency with the same frequency difference between Pi and g-ATP (Fig. 10). This decrease in the intensity of Pi (DMPi) is then used to determine VATP. Because g-ATP is saturated the term k1MATP(t) (Eq. 7) is set to zero simplifying the equation. When integrated and applied to experimental conditions of long saturation times we have: M Pi = 1 (8) M 0 Pi 1 + k-¢1T1Pi Knowing that the T1Pi obtained under g-ATP saturation (T ¢1Pi) is given by 1 / T1¢ Pi = 1 / T1Pi + k-¢1 , Eq. 8 simplifies to: k-¢1 = (1 - M Pi / M 0 Pi )

(9)

T1¢ Pi Therefore, as seen in Eq. 9, the decrease in the intensity of Pi is directly proportional to VATP. 1.4. 13C MRS/13C NMR

13

C MRS has been applied in several studies of intermediary metabolism since late 1970s (20, 21). Due to the low natural abundance of the carbon 13 isotope, such studies have mostly focused on the most abundant metabolites, particularly lipids (22) and glycogen (23). An alternative approach has been implemented for increasing 13 C enrichments considerably above natural abundance levels. By means of the administration of 13C-enriched substrates (24, 25), 13 C enrichments of several metabolic intermediates are increased to levels compatible with adequate detection by 13C MRS and a multitude of studies were made possible by in vivo 13C MRS (for a review see Bachelard et al. (26)). Analysis of Krebs cycle activity is frequently made by the detection of 13C incorporation in metabolic intermediates like glutamate. Most metabolic analyses make use of 13C fractional enrichments due to resolution constraints (14, 27, 28) but in some circumstances richer positional enrichment

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(isotopomer) information can be derived from the analyses of 13C NMR spectra both in vivo (29), ex vivo (30) and in vitro (11, 31). The ability to discriminate between isotopomers is crucial for analysis of complex metabolic pathways and for determining the preferences of exogenously provided substrates under particular pathophysiological conditions (32, 33). 13C NMR is also a powerful tool for detecting alterations in Krebs cycle and transaminase activities (11). Changes in these fluxes reflect themselves in major alterations in the incorporation of 13C in TCA cycle intermediates and in metabolites in exchange with them. Kinetic measurement of 13C incorporation in glutamate constitutes the ultimate tool for analyzing metabolite trafficking between mitochondria and cytosol (11, 34). Figure 11 shows two 13C NMR spectra from hearts perfused for 30 min with [3-13C]lactate plus [3-13C]pyruvate. Under most circumstances, the NMR spectrum of heart tissue is dominated by the resonances due to glutamate (Fig. 11a), being resonances due to other metabolic intermediates too low to be detected. However, changes in the cytosolic redox, caused by an inversion of the physiologic ratios of lactate/pyruvate (from 10:1 to a 1:10), cause major redistributions in the intermediate metabolite pools and other metabolites such as citrate, fumarate, malate, and aspartate appear with high intensity (Fig. 11b) (35). This allows direct observation of isotopomers due to metabolic intermediates within the Krebs cycle (citrate, fumarate, and malate) and a comparison with isotopomers from aminoacids (glutamate and aspartate) in exchange with them. A direct evaluation of 13C labeling of Krebs cycle intermediates is crucial for measuring Krebs cycle flux without the need to assume fast exchange involving transamination reactions (14, 36). In most circumstances, however, the levels of Krebs cycle intermediates are too small to be detected by direct detection of 13C incorporation by 13C NMR. Indirect detection NMR methodologies, like J-Res-HSQC (37) and HMQC-TCOSY (38), have been proven very valuable in allowing the reduction of the detection limit but detection limits are still too high to allow wide application. 1.5. How to Overcome 13 C NMR Sensitivity Problems?

NMR is an inherently low sensitive technique. According to the Boltzmann distribution, a sample at 37°C in a magnetic field of 9.4 T has a spin difference between the low and upper state, i.e., nuclear polarization, of only 31 spins. In particular, the low gyromagnetic ratio of 13C limits the use of this stable isotope to the analysis of highly concentrated metabolites. Several methods, known as hyperpolarization, have therefore been proposed to increase the nuclear polarization of a sample. Such methods include optical pumping (39), para-hydrogen-induced polarization (PHIP) (40, 41), and dynamic nuclear polarization (DNP) (42, 43). In this section, the general principles of DNP are explored.

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a

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Fig. 11. 11.7 T 13C NMR spectra of extracts from rat hearts perfused with 0.4 mM [3-13C] pyruvate plus 4.0 mM [3-13C]lactate (a) or 4.0 mM [3-13C]pyruvate plus 0.4 mM [3-13C] lactate (b). Aside from glutamate carbons (Glu-C2, Glu-C3, and Glu-C4), expansions in b also show (left to right ) citrate C3 (Cit-C3), malate C2 (Mal-C2), aspartate C2 (Asp-C2), citrate C2 and C4 (Cit-C2C4), malate C3 (Mal-C3), and aspartate C3 (Asp-C3). Expansions in b refer to glutamate carbons C2 (Glu-C2), C3 (Glu-C3), and C4 (Glu-C4). 1.5.1. Dynamic Nuclear Polarization

During an experiment of DNP, the sample, placed in a diamagnetic insulator with a small concentration of paramagnetic substance with a Larmor frequency we, is irradiated with a microwave frequency leading to enhanced polarization. For maximum polarization this is kept a temperature close to 0 K. To better understand this mechanism, consider a sample with totally unpolarized nuclear spins I = 1/2 of Larmor frequency wI mixed with a paramagnetic substance with totally polarized electron spins S = 1/2 of Larmor frequency wS coupled to the nuclear spins by dipolar interactions (Fig. 12a). In the presence of B0, the lower energy state is populated

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Fig. 12. Mechanism of dynamic nuclear polarization (DNP) by which an unpolarized sample can be polarized in the presence of paramagnetic substances. In an unperturbed state, in the presence of an external magnetic field (B0) the nuclear spins I are equally distributed among the two energy levels and the electron spin S are coupled to spins S (a). In the presence of a microwave of frequency W = wS − wI , the reorientation of the spins S would prompt a change in the nuclear spin I in a way that spins S and I are paralleled (b). Because the relaxation rate of spins S is much faster than spins I, the reorientation of spin S (W = 0), is not followed by a reorientation of spins I leading to an increase of the net magnetization. If instead W = wS + wI , the polarization is inverted (d–f).

with nuclear spins paralleled with B0 while in the upper state the spins are reversed. In order to have a stable coupling between spins S and I, the electron spins are antiparalelled with respect to B0. To change the orientation of the spins, radiation in the range of microwave must be supplied. If this system is irradiated with an external source of energy W, such that W = wS − wI, assuming that DwS << wI, a spin I antiparallel with respect to B0 is excited reversing its direction. At the same time orientation of the spin S is also altered (Fig. 12b). At this stage, the rate of relaxation of each spin, i.e., how fast the spins return to the original orientation, is crucial to allow the enhancement of nuclear polarization. If the relaxation rates for S and I were similar, the final orientation of the spins would be the same as in Fig. 12a. However, while for the spins S this relaxation constant is »103/s, it is much slower for the spins I (»10−3/s) allowing the number of parallel spins I to increase, enhancing therefore the polarization (Fig. 12c) (42). It is easy to see that if W = wS + wI, the polarization is inverted (Fig. 12d–f ).

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The lasting effect of this polarization enhancement is very important. For instance, while hyperpolarized 13C has a T1 of 10,800 s, at 1.5 K, it suffers a decrease of ~96% to 435 s at 10 K (43). This imposes an important limitation on the use of hyperpolarized metabolites as tracers to follow metabolism in vivo. However, a recent method was developed to hyperpolarize organic molecules in liquid state (44). Using this approach, the organic tracer is polarized in the solid state and quickly dissolved conserving the high polarization for ~60 s. Despite the short T1, relative to what can be obtained at temperatures <4 K, this allows a signal-to-noise ratio that is several orders of magnitude higher than the one obtained with unpolarized tracers (44). The ability to increase the signal-to-noise ratio, characteristically low in 13C NMR spectra, is a major breakthrough in the study of metabolism in vivo. This increased sensitivity was initially used in the imaging of tumors and perfusion mapping (45). However, it has now been used to measure the flux through a single enzyme-catalyzed step. In an experiment performed by Malloy et al., the infusion of hyperpolarized [1-13C]pyruvate was used to measure the flux through PDH in perfused hearts (46). In this experiment, the spectra, dominated by the resonance of 13C in C1 pyruvate and with a time resolution of 1 s, showed the timecourse of enrichment of [1-13C]pyruvate, [1-13C]lactate, and the 13 CO2 produced by the oxidation of pyruvate. Using this approach, it was also possible to observe the inhibitory effects of fatty acid oxidation on PDH since no production of H13CO3− was observed. In a different study, the same authors were able to detect and quantify a delay in the activation of the flux through PDH in ischemic hearts following reperfusion (47). In such experiment, hearts subjected to a 10-min ischemia were reperfused with hyperpolarized [1-13C]pyruvate. It was observed that during the first 90 s of reperfusion no H13CO3− was detected suggesting total inhibition of PDH and redirection of the metabolic flux toward [1-13C]lactate synthesis. Only after 15 min of reperfusion, the activation of PDH was observed. The use of hyperpolarized [1-13C]pyruvate is specially relevant due to its high T1 relaxation time (40–50 s). As mentioned above, the high signal intensity of a hyperpolarized 13C lasts for a very short period of time. Therefore, a long T1 as the one observed for [1-13C]pyruvate allows the detection of other enriched metabolites that resulted from subsequent reactions. It is the case, for instance, of the study performed by Tyler et al. (48). In this experiment, [2-13C]pyruvate was infused in hearts and, also with a time resolution of 1 s, the time course of enrichments of pyruvate, lactate, acetylcarnitine, citrate, and glutamate was followed. The ability to follow the enrichments of these metabolites is of extreme importance since it allows the direct measure of the in vivo fluxes through PDH and TCA cycle giving important information about the metabolic

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profile of a tissue in a wide range of physiological and pathophysiological settings. Other tracers have been used to quantify mitochondrial metabolism. Lerche et al., used hyperpolarized [1-13C] acetate to measure, in anesthetized animals, the flux through acetyl-CoA synthetase and acetylcarnitine transferase in heart and liver (49). Although this is a major breakthrough in the study of metabolism the use of hyperpolarized substrates is still a challenge and requires further study and optimization. The level of enrichment that can be detected in a 1-s resolution is again limited to metabolites that exist in high concentrations in the cells. Therefore, the development of tracers that are able to retain high polarization is key in the further development of this field. In conclusion, NMR in its multiple possible applications (conventional high-resolution NMR of tissue extracts, high rotation magic angle spinning (HRMAS) of tissue biopsies and in vivo MRS), constitutes a “unique tool” for evaluating the bioenergetic status of a whole organism or an isolated perfused organ, providing unique information on the status of the metabolic machinery and allowing the monitoring of pathophysiological conditions through the detection of modifications in the relative and absolute levels of metabolic intermediates and through the analysis of the alterations in the rates of stable isotope incorporation. Multiple approaches can be considered toward an accurate evaluation of mitochondrial performance and their choice is multifactorial. Ultimately, all technological developments aim at determining metabolic fluxes in humans in vivo (50, 51) and improvements in MRS techniques (52, 53) assume special relevance toward such goal.

2. Materials 2.1. Isolated Heart Perfusion

Isolated rat hearts are perfused in Krebs–Henseleit (KH) bicarbonate buffer supplemented with 13C-enriched metabolites. 1. KH bicarbonate buffer: 119.2 mM NaCl, 4.7 mM KCl, 1.2 mM CaCl2, 1.2 mM MgSO4, and 25 mM NaHCO3. The buffer is saturated with 95% O2–5% CO2, maintaining a pH of 7.4. All salts for media preparation were obtained from SigmaAldrich (Sintra, Portugal). 2. Typical 13C-enriched metabolites include: 1.2 mM [3-13C] lactate, 0.09 mM [3-13C]pyruvate, 0.05 mM [1,3-13C2] acetoacetate, 0.12 mM [1,3-13C2] b-hydroxybutyrate, from Cambridge Isotope Laboratories (Andover, MA) and 0.35 mM [U-13C]long-chain fatty acids (LCFAs), from Isotec Inc. (Miamisburg, OH). Glucose is also frequently present at a concentration of 5.5 mM but in most cases is provided at natural abundance levels since its contribution to acetylCoA is significantly smaller than from other sources.

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3. Other mixtures of 13C-enriched metabolites can be used in circumstances of significant modification in substrate preference by the heart. One of those mixtures might involve the switch between [U-13C]LCFAs and [U-13C]glucose. 2.2. Hippocampal Slice Superfusion

Hippocampi are rapidly removed from freshly isolated rat brain and transversely cut in 400 mm slices using a McIlwain tissue chopper. Superfusion media: 115 mM NaCl, 25 mM NaHCO3, 3 mM KCl, 1.2 mM KH2PO4, 2 mM CaCl2, 1.2 mM MgSO4, 5.5 mM glucose, and 2 mM sodium acetate, continuously gassed with carbogen, pH 7.4. Glucose and acetate are either provided at natural abundance enrichment levels – period of slice stabilization – or enriched in carbon 13, [U-13C]glucose and [2-13C]acetate, during the isotopic labeling period. In order to reduce the time needed to reach isotopic and metabolic steady state, the medium is supplemented with 50 mM 4-aminopiridine, to trigger intermittent burst-like slice stimulation.

2.3. Samples for High-Resolution NMR

Lyophilized extracts are redissolved in 99.9% D2O and the pD adjusted to ~7.0 with DCl or NaOD solutions (Sigma-Aldrich, Sintra, Portugal). In most circumstances, proton NMR spectra are acquired for quantitative purposes and a suitable internal standard is added to the NMR sample for absolute measures of intermediary metabolite levels. We have been using sodium fumarate (SigmaAldrich, Sintra, Portugal), since this metabolite resonates at 6.5 ppm, away from the crowded regions of the proton NMR spectrum and is present in tissues in concentrations too low to be of any significance in terms of its contribution to the added quantity (~2 mM final concentration in NMR sample).

3. Methods 3.1. Perchloric Acid Extraction

Perchloric acid extraction for preparation of tissue extracts for high-resolution NMR analysis. 1. Deep-freezed tissue is pulverized into a fine powder using nitrogen precooled mortar and pestle. 2. Two Volumes of ice-cold perchloric acid (PCA) 7% are added for each gram of wet tissue; upon addition of PCA vortex and allow tissue to melt – keep tissue in ice. Perform vigorous vortex several times for sample homogenization. 3. Centrifuge the homogenate at 2,520 g, at 4°C, during 15 min. The supernatant (aqueous phase) containing water-soluble metabolites is recovered and the pellet containing membranes, cell debris, and other metabolites discarded.

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4. Neutralize the supernatant with KOH (pH 6.9–7.0) keeping the solution in ice. This ensures a more significant precipitation of the salt being formed, potassium perchlorate (KClO4). 5. Centrifuge at 2,520 g, at 4°C, during 15 min to remove precipitated salt. Resulting supernatant is subject to lyophilization to concentrate intermediate metabolites. 6. In the PCA extract all water-soluble metabolites can be found including aminoacids (e.g., glutamate, glutamine, aspartate, alanine), ketoacids (e.g., pyruvate, oxaloacetate, a-ketoglutarate), lactate, glucose. 7. The obtained extract still contains significant amount of salts. This could constitute a major problem when acquiring highresolution NMR spectra. Two further steps can help improve the spectra quality. First, upon dissolution of the extract in 2 H2O allow the sample to stay at 4°C. This will cause further precipitation of salt (mostly KClO4) and sample viscosity will be significantly reduced. Second, add a considerable amount of a chelating agent, namely ethylenodiaminotetracetic acid (disodium salt), to complex most cations in solution which otherwise will cause significant line broadening in the NMR spectrum. 8. While performing this extraction procedure special care has to be taken while adjusting the pH with KOH (step 4). This adjustment to pH ~7.0 has to be made slowly since an incorrect adjustment of the pH can lead to destruction of the sample upon liophilization if pH is left considerably low, or to an excessive increment in solution salt content due to the need to add further PCA if pH keeps going above 7.0 by excessive addition of KOH. Several solutions of KOH must be prepared in order to allow the smallest addition of KOH solution but at the same time be as correct as possible in pH adjustment. In the initial steps, a saturated solution of KOH should be used but as pH approaches the target value KOH solutions 0.5 M or even 0.1 M should be used. At all times try to keep the added volume to a minimum since an increment in sample solution will consequently increase the amount of salt which is dissolved and is not removed by the second centrifugation step. Still associated with pH misadjustment is another problem which is frequently encountered when running NMR spectra from tissue extracts. If the pH on the dissolved lyophilized sample approaches the pKa value of the amino group of aminoacids like glutamate, then due to chemical exchange phenomena the resonances of those molecules suffer considerable line broadening. This effect can be avoided by dissolving the tissue extract using 2H2O containing a significant buffer capacity, given for example by a phosphate buffer.

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1. While this might sound a simple matter, and in fact it is, one frequently come across incorrect procedures in an NMR laboratory setting which must be avoided if the data is intended to have any meaning. (notes). The first worry should always be to have the “best sample” the possible. This implies careful sample handling to avoid several types of interferences: (a) avoid high salt content in samples whenever possible; (b) use adequate (see specifications by the NMR spectrometer provider) sample heights and keep them constant in all analysis – this simple caution speeds up considerably NMR analysis by avoiding the need for significant NMR field adjustments on sample change; (c) avoid by all means the presence of air bubbles in the sample – this causes abnormal resonance broadening. 2. Most one-dimensional (1D) proton NMR spectra used in metabolic studies are acquired with the intent of quantifying metabolites in solution. In order to be quantitative, fully relaxed spectra need to be obtained. This implies correct knowledge about the relaxation behavior of all molecules in the solution being analyzed. This does not necessarily imply that a full relaxation (T1/T2) analysis has to be made for each sample but in the beginning of any study an estimation of those parameters should be made to warrant that spectra will be acquired quantitatively. 3. The simplest 1D NMR experiment possesses three essential stages: (a) a first stage which comprises a delay time, which needs to be adjusted in accordance with the relaxation behavior of the sample. This stage could also include a period of solvent saturation in case the solvent resonance is much more intense than the resonances of the metabolites being analyzed; (b) a second stage which is simply the excitation period and implies the application of a radiofrequency pulse for a given duration and power. This again has to be adjusted according to sample specifications; high-salt content leads to significant increases in pulse duration and pulse calibrations should be made prior to quantitative spectra acquisition; (c) a third stage which consists in free-induction-decay (FID) acquisition – the duration of this period, also called acquisition time (at) is chosen to provide enough signal digitization to derive well-resolved spectra. The total duration of the three stages is frequently referred to as the experimental “repetition time” (RT). 4. Adequate signal-to-noise ratio is required for accurate determinations of metabolite concentrations. Increment in signalto-noise is achieved by increasing the number of transients accumulated for each FID. In the case of tissue extracts, there are some metabolites which are frequently abundant such as glutamate and lactate and for which there is not much need for

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transient accumulation for signal averaging. However, for metabolites less abundant like many of the Krebs cycle intermediates, there is frequently the need for performing signal averaging and total acquisition times can easily be 30 min to several hours, depending on the amount of tissue material available. The RT used between transients is a function of the relaxation behavior as stated above but as well a function of the pulse width being used. Increased signal-to-noise ratios are possible maintaining the quantitative character of the spectrum by lowering considerably the excitation pulse length from a maximum of 90°. Frequently used parameters for tissue PCA extracts include a radiofrequency pulse of 30° and a RT of 15 s, from which 9 s are the relaxation delay, 3 s are the presaturation period for solvent suppression, using a saturation pulse with the lowest possible saturation power to avoid saturation of nearby resonances, and finally 3 s of acquisition time for FID accumulation. 5. In summary, a quantitative NMR spectrum requires long enough RT to ensure full relaxation and good signal-to-noise ratio to allow accurate estimates of metabolite concentrations by signal deconvolution and comparison with a well-defined internal standard. 3.3. Acquisition of High-Resolution 13 C NMR Spectra

1. All aspects mentioned above apply to 13C NMR in the same way as described for 1H NMR. However, while using 1H NMR one can be absolutely quantitative, when using 13C NMR spectra several extra considerations need to be taken into account in order to obtain acceptable quantitative information, namely nuclear Overhauser effect (NOE) and signal saturation by oversampling. 2.

13

C NMR spectra are almost always acquired with 1H broadband decoupling to remove scalar coupling from directly coupled protons and from protons attached to neighboring carbons. The need for broadband decoupling arises from the need to reduce spectral complexity and, most importantly, to improve considerably the signal-to-noise of 13C NMR spectra. This improvement in signal-to-noise can be further enhanced by allowing for NOE to build up during the delay before radiofrequency excitation. While this NOE improves signal-tonoise and allows a reduction on detection limits, it confuses quantification procedures due to the fact that NOE effects are not identical for all aliphatic carbons and are essentially absent in quaternary carbons. The only possibility for being quantitative resides then in finding the NOE factors so that normalization of signal intensities can be made after spectrum acquisition and quantification.

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3. The problem of oversampling is always present when acquiring 13 C NMR spectra of tissue extracts. This is the result of a significant difference between the relaxation behaviors of aliphatic carbons relative to quaternaries. Differences in relaxation times of one order of magnitude are frequently found and allowing full relaxation of quaternary carbons would essentially render the experiment obsolete due to total lack of signal-to-noise. Most frequently enough time is given to allow full relaxation to all aliphatic carbons of a given metabolite and quantitative comparisons can be made between them but such time is too small to allow full relaxation of carboxyl or carbonyl carbons. The practical result is a 13C NMR spectrum in which the aliphatic carbons appear much more intense than the quaternaries and direct comparison of their areas is totally impossible. 4. A typical 13C NMR spectrum of a tissue extract uses a repetition time between transients of 3 s and a radiofrequency pulse width equivalent to 45°. These parameters ensure an almost complete relaxation of aliphatic carbons and, providing that adequate NOE normalization factors are obtained for such samples, the areas derived from the 13C NMR spectrum can be used as a measure of 13C enrichment. 3.4. Isolated Heart Perfusion

1. Heart from mice or rats can be perfused after removal from the animal using the Langendorff or retrograde heart perfusion. 2. After general anesthesia mice or rat hearts are rapidly removed and placed in ice cold perfusion medium, the aorta is immediately cannulated, and hearts subject to a perfusate column height of 100 cm H2O. The excess tissue and fat can be removed while heart rate stabilizes. The modified Krebs–Henseleit buffer, containing 119.2 mM NaCl, 4.7 mM KCl, 1.25 mM CaCl2, 1.2 mM MgSO4, and 25 mM NaHCO3, is continuously bubbled with 95% O2–5% CO2 and in most studies is not recirculated. 3. Several metabolic analyses can be performed using 13C-enriched substrates as a supplement of the KH bicarbonate buffer. 4. In substrate competition measurements by 13C NMR isotopomer analysis, four distinct sources of acetyl-CoA can be distinguished in a single experiment. Most frequently the mixture chosen contains [U-13C]LCFA, [3-13C]lactate plus [3-13C]pyruvate, [1,3-13C]acetoacetate plus [1,3-13C]b-hydroxybutyrate, and glucose at natural abundance enrichment levels. These four varieties of substrates originate acetyl-CoA with alternate 13 C-labeling patterns which are distinguishable by analysis of the 13C labeling patterns of glutamate carbons C4 and C5. This substrate competition analysis does not require metabolic and isotopic steady state to be reached. Simple equations correlate the 13C multiplets in glutamate carbons with the composition of the acetyl-CoA pool at any time of the perfusion.

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5. In evaluations of Krebs cycle kinetics most frequently only one substrate is provided with 13C enrichment. The choice of the substrate is made so that the acetyl-CoA generated becomes enriched in carbon 2 of the acetyl moiety. This will enrich carbon 4 on a first turn of the cycle and that labeling will appear in carbons C2 or C3 on a second turn and eventually in carbon 1 in subsequent turns. The appearance of multiply enriched glutamate will thus be a measure of Krebs cycle turnover and could be correlated with measures of oxygen consumption. The choice of singly labeled precursors capable of forming [2-13C]acetyl-CoA is also crucial from a sensitivity point of view. If the enrichment is to be followed ex vivo, having the heart being perfused inside the magnet, than a smaller multiplicity will be favorable for detection by direct 13C MRS and better time resolution could be achieved in the analysis. 6. The entire all-glass perfusion system is jacketed and maintained at 37°C throughout the entire duration of the perfusion. 7. The perfusion system can have several buffer chambers but typically two chambers are considered. In one of the chambers there is buffer with unenriched substrates, which is used for initial washing and stabilization procedures and a second chamber containing the 13C-enriched substrates which is used for metabolic studies by 13C NMR isotopomer analysis. Of course such chambers can also be used to follow the effects on metabolism due to some hormones and or metabolic inhibitors. 8. By the end of the perfusion protocol the perfused heart is in most cases freeze-clamped using nitrogen precooled aluminum tongs. The obtained tissue is ready for extraction protocols, namely PCA extraction as described above. 3.5. In Vivo 1H NMR Spectroscopy

1. High-resolution spectra of tissue extracts are acquired using NMR tubes containing an homogeneous tissue extract and vertical narrow bore high-magnetic field spectrometers. In these circumstances, it is quite simple to achieve excellent field homogeneity and NMR spectra are of very-high resolution. However, when acquiring in vivo 1H NMR spectra the conditions are significantly altered and achieving excellent field homogeneity in an heterogeneous sample using a horizontal wide bore magnet becomes very demanding. 2. The object of analysis (e.g., brain, muscle) should be positioned in the isocenter of the magnet and fast-spin-echo images with repetition time of 5 s, echo time of 52 ms, and echo train length of 8 are used to identify the substructure of interest (e.g., hippocampus) based on anatomical landmarks. 3. Shimming for field homogeneization can be performed using specific programs, like FAST(EST)MAP (54), and 1H NMR spectra are subsequently acquired from the volume of interest

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(VOI) 18 mL placed in the left hippocampus. A sequencedesigned SPECIAL with echo time of 2.8 ms and repetition time of 4 s (55) can be used for localization. 4. Spectral analysis of in vivo 1H NMR spectra can be carried out using LCModel (55) including a macromolecule spectrum in the database (35). The unsuppressed water signal measured from the same VOI can be used as an internal reference for the quantification of several metabolites, namely glucose, ascorbate, phosphorylethanolamine, creatine, phosphocreatine, myo-inositol, taurine, N-acetylaspartate, aspartate, glutamate (Glu), glutamine, GABA, alanine, lactate, b-hydroxybutyrate, glycerophosphorylcholine, phosphorylcholine, GSH, N-acetylaspartylglutamate, and scyllo-inositol. The Crame´r-Rao lower bound provided by LCModel can be used as a measure of the reliability of the apparent metabolite concentration quantification (56). Metabolite concentrations with Crame´r-Rao lower bound higher than 25% should not be included in the analysis. Spectral quality is evaluated by analyzing the metabolite line widths and signal-to-noise ratio that were provided by LCModel. 3.6. Hyperpolarized 13 C NMR Spectroscopy in the Isolated Perfused Heart

1. In order to improve signal-to-noise by several orders of magnitude hyperpolarized [1-13C]pyruvate can be used as a substrate for the isolated perfused rat heart. Hyperpolarization can be achieved using procedures similar to those described by Golman et al. (57) or Merritt et al. (46, 47). 2. The polarization process is started approximately 90 min before each experiment. An aliquot, ~20 mL, of a 3-M solution of [1-13C] pyruvate, prepared in deionized H2O is mixed with an equal volume of 16.6 mM trityl radical in 50/50 glycerol/water. 3. The solutions can be polarized using a commercial device (HyperSense™, Oxford Instruments Molecular Biotools, Abingdon, UK) to a level of ~15% after 90 min of microwave irradiation at 1.4°K. 4. The polarized sample is rapidly thawed by dissolution in 4 mL of heated water containing 0.85 mM Na2EDTA. 5. 3 mL of the resulting solution are further diluted into 20 mL of substrate-free perfusate to achieve a final concentration of 2 mM hyperpolarized [1-13C]pyruvate. 6. All the above steps have to be performed very rapidly. Total time for dissolution, ejection, and further dilution should not exceed ~10 s. 7. The solution can be injected gently and continuously by catheter into the perfusion column directly above the heart over a period of 90 s. The temperature of all solutions entering the heart has to be kept at 37°C and the pH kept between 7.3 and 7.4. The NMR console should be triggered to start acquisition at the end of the dissolution process.

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4. Notes In studies of intermediary metabolism, both in vivo and in vitro, which apply NMR methodologies as the technique of choice for determining the levels and enrichment of intermediary metabolites major caution should be taken in the following: (1) resolution of NMR spectra should be the best at all times to allow proper separation of all resonances in the sample of interest. Only wellresolved spectra are possible to use for adequate quantification of intermediary metabolites present; this is certainly user driven but nowadays there are many mechanisms which allow automatization of the procedure so that sufficient quality can be reached at all times; (2) fully relaxed spectra should be acquired whenever possible – partially saturated spectra can be used in some circumstances providing that adequate knowledge of the system exists and correction factors had been properly derived; (3) when using isotopically enriched substrates for monitoring specific metabolic pathways the correct choice of such metabolites might be the difference between a successful experience or a total failure. Some mathematical models exist which allow simulation of metabolic pathways – these can be used to probe for the perfect “cocktail” of isotopicaly enriched substrates. This is of the utmost importance for two major reasons. First, only some tracers are able to provide the metabolic information being other unspecific. Second, the metabolite needs to be considerably metabolized by the tissue so that enough enrichment is attained in intermediary metabolites compatible with their detection in acceptable acquisition times in the spectrometer. As an example, one should not use 13C-enriched glucose for monitoring Krebs cycle kinetics when using isolated hearts from control animals since these prefer LCFAs as the source for acetyl-CoA. Thus, a prior knowledge about substrate preferences/ metabolic routes has to be included in the design of the experimental approach. References 1. Kumar R, Ernst RR, Wuthrich K (1980) A two-dimensional nuclear overhauser enhancement (2D NOE) experiment for the elucidation of complete proton cross-relaxation networks in biological macromolecules. Biochem Biophys Res Commun 95:1–6 2. Marion D, Wuthrich K (1983) Application of phase sensitive two-dimensional correlated spectroscopy (COSY) for measurements of H-1H-1 spin-spin coupling-constants in proteins. Biochem Biophys Res Commun 113:967–974

3. Hoult DI, Busby SJ, Gadian DG, Radda GK, Richards RE, Seeley PJ (1974) Observation of tissue metabolites using 31P nuclear magnetic resonance. Nature 252:285–287 4. Wallace DC (2007) Why do we still have a maternally inherited mitochondrial DNA? Insights from evolutionary medicine. Annu Rev Biochem 76:781–821 5. Maassen JA, Janssen GM, t Hart LM (2005) Molecular mechanisms of mitochondrial diabetes (MIDD). Ann Med 37:213–221

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6. Wallace DC, Fan W (2010) Energetics, epigenetics, mitochondrial genetics. Mitochondrion 10:12–31 7. Szendroedi J, Roden M (2008) Mitochondrial fitness and insulin sensitivity in humans. Diabetologia 51:2155–2167 8. Petersen KF, Dufour S, Befroy D, Garcia R, Shulman GI (2004) Impaired mitochondrial activity in the insulin-resistant offspring of patients with type 2 diabetes. N Engl J Med 350:664–671 9. Chance EM, Seeholzer SH, Kobayashi K, Williamson JR (1983) Mathematical analysis of isotope labeling in the citric acid cycle with applications to 13C NMR studies in perfused rat hearts. J Biol Chem 258:13785–13794 10. Weiss RG, Gloth ST, Kalil-Filho R, Chacko VP, Stern MD, Gerstenblith G (1992) Indexing tricarboxylic acid cycle flux in intact hearts by carbon-13 nuclear magnetic resonance. Circ Res 70:392–408 11. Carvalho RA, Rodrigues TB, Zhao P, Jeffrey FM, Malloy CR, Sherry AD (2004) A 13C isotopomer kinetic analysis of cardiac metabolism: influence of altered cytosolic redox and [Ca2+]. Am J Physiol Heart Circ Physiol 287(2): H889–H895 12. Fitzpatrick SM, Hetherington HP, Behar KL, Shulman RG (1990) The flux from glucose to glutamate in the rat brain in vivo as determined by 1H-observed, 13C-edited NMR spectroscopy. J Cereb Blood Flow Metab 10(2):170–179 13. Doan BT, Autret G, Mispelter J, Méric P, Même W, Montécot-Dubourg C, Corrèze JL, Szeremeta F, Gillet B, Beloeil JC (2009) Simultaneous two-voxel localized 1H-observed 13 C-edited spectroscopy for in vivo MRS on rat brain at 9.4T: Application to the investigation of excitotoxic lesions. J Magn Reson 198:94–104 14. Mason GF, Gruetter R, Rothman DL, Behar KL, Shulman RG, Novotny EJ (1995) Simultaneous determination of the rates of the TCA cycle, glucose utilization, alpha-ketoglutarate/glutamate exchange, and glutamine synthesis in human brain by NMR. J Cereb Blood Flow Metab 15:12–25 15. Gruetter R, Seaquist ER, Ugurbil K (2001) A mathematical model of compartmentalized neurotransmitter metabolism in the human brain. Am J Physiol Endocrinol Metab 281:E100–E112 16. Brindle KM, Blackledge MJ, Challiss RA, Radda GK (1989) 31P NMR magnetizationtransfer measurements of ATP turnover during steady-state isometric muscle contraction in the rat hind limb in vivo. Biochemistry 28: 4887–4893

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41. Haake M, Natterer J, Bargon J (1996) Efficient NMR pulse sequences to transfer the parahydrogen-induced polarization to hetero nuclei. J Am Chem Soc 118:8688–8691 42. Abragam A, Goldman M (1978) Principles of dynamic nuclear-polarization. Rep Prog Phys 41:395–467 43. Wolber J, Ellner F, Fridlund B, Gram A, Johannesson H, Hansson G, Hansson LH, Lerche MH, Mansson S, Servin R et al (2004) Generating highly polarized nuclear spins in solution using dynamic nuclear polarization. Nuc Inst Methods Phys Res A 526:173–181 44. Ardenkjaer-Larsen JH, Fridlund B, Gram A, Hansson G, Hansson L, Lerche MH, Servin R, Thaning M, Golman K (2003) Increase in signal-to-noise ratio of >10,000 times in liquidstate NMR. Proc Natl Acad Sci USA 100:10158–10163 45. Aime S, Dastru W, Gobetto R, Santelia D, Viale A (2008) Agents for polarization enhancement in MRI. Handb Exp Pharmacol 185(pt 1): 247–272 46. Merritt ME, Harrison C, Storey C, Jeffrey FM, Sherry AD, Malloy CR (2007) Hyperpolarized 13 C allows a direct measure of flux through a single enzyme-catalyzed step by NMR. Proc Natl Acad Sci USA 104:19773–19777 47. Merritt ME, Harrison C, Storey C, Sherry AD, Malloy CR (2008) Inhibition of carbohydrate oxidation during the first minute of reperfusion after brief ischemia: NMR detection of hyperpolarized 13CO2 and H13CO3. Magn Reson Med 60:1029–1036 48. Schroeder MA, Atherton HJ, Ball DR, Cole MA, Heather LC, Griffin JL, Clarke K, Radda GK, Tyler DJ (2009) Real-time assessment of Krebs cycle metabolism using hyperpolarized 13 C magnetic resonance spectroscopy. FASEB J 23:2529–2538 49. Jensen PR, Peitersen T, Karlsson M, In’t Zandt R, Gisselsson A, Hansson G, Meier S, Lerche MH (2009) Tissue-specific short chain fatty acid metabolism and slow metabolic recovery after ischemia from hyperpolarized NMR in vivo. J Biol Chem 284:36077–36082 50. Tkác I, Oz G, Adriany G, Uğurbil K, Gruetter R (2009) In vivo 1H NMR spectroscopy of the human brain at high magnetic fields: metabolite quantification at 4T vs. 7T. Magn Reson Med 62:868–879 51. Sailasuta N, Robertson LW, Harris KC, Gropman AL, Allen PS, Ross BD (2008) Clinical NOE 13C MRS for neuropsychiatric disorders of the frontal lobe. J Magn Reson 195:219–225 52. Henry PG, Adriany G, Deelchand D, Gruetter R, Marjanska M, Oz G, Seaquist ER, Shestov A,

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Chapter 19 Computational Modeling of Mitochondrial Function Sonia Cortassa and Miguel A. Aon Abstract The advent of techniques with the ability to scan massive changes in cellular makeup (genomics, proteomics, etc.) has revealed the compelling need for analytical methods to interpret and make sense of those changes. Computational models built on sound physico-chemical mechanistic basis are unavoidable at the time of integrating, interpreting, and simulating high-throughput experimental data. Another powerful role of computational models is predicting new behavior provided they are adequately validated. Mitochondrial energy transduction has been traditionally studied with thermodynamic models. More recently, kinetic or thermo-kinetic models have been proposed, leading the path toward an understanding of the control and regulation of mitochondrial energy metabolism and its interaction with cytoplasmic and other compartments. In this work, we outline the methods, step-by-step, that should be followed to build a computational model of mitochondrial energetics in isolation or integrated to a network of cellular processes. Depending on the question addressed by the modeler, the methodology explained herein can be applied with different levels of detail, from the mitochondrial energy producing machinery in a network of cellular processes to the dynamics of a single enzyme during its catalytic cycle. Key words: Systems biology, Kinetic and thermo-kinetic models, Mitochondrial energy transduction, Ordinary differential equations, Model parameters

1. Introduction The emergence of “Systems Biology” signals the recognition of an integrative era in Biology, by contrast to the analytical period of biochemistry, cellular, and molecular biology (1–4). Systems biology encompasses a series of integrative approaches attempting to describe extensive changes in gene expression at the RNA, protein, or in the metabolite cellular makeup. In this regard, Systems Biology combines high-throughput technologies with a variety of quantitative methods, including computational modeling,

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0_19, © Springer Science+Business Media, LLC 2012

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in order to gain a more comprehensive understanding of systemic changes (5). Computational approaches include thermodynamic, kinetic, and stoichiometric models. In the past, thermodynamic models have been extensively used for describing energy transduction in microbial physiology and mitochondrial energetics (6–10). The advantage of these earlier attempts was their simplicity and agreement with the fundamental principles ruling biological free energy transduction, i.e., the thermodynamic laws. However, the lack of mechanistic details accounting for many of the observed behaviors was a serious drawback. A kinetic approach allows mechanisms and regulatory properties to be specified. A direct link between the purely thermodynamic approach and the kinetic one described in this work, is that at steady state, and assuming that fluxes are linearly related with their respective driving forces, kinetic information is contained implicitly within the phenomenological coefficients in the Onsager equation (11). Stoichiometric models can encompass large networks of biological processes, taking into account only the net stoichiometry of the transformations occurring during their operation (12–14). They will render a space of solutions for the system and as no explicit regulatory interactions are accounted for in this formulation, they will inform essentially about steady-state fluxes without any possibility of transient behavior or regulatory interactions. Kinetic models can circumvent the limitations of both thermodynamic and stoichiometric, approaches since their description comprises regulatory interactions and kinetic mechanisms (15, 16). Limitations in the availability of quantitative kinetic information (e.g., maximal velocities, affinity constants) are among the drawbacks of the kinetic approach. However, when quantitative data is lacking, the model can be used as a tool for testing hypotheses concerning the mechanism(s) underlying an observed behavior. Kinetic models can also be phenomenological, i.e., not based on known or hypothetical chemical or transport reaction schemes. The power-law formalism (17) is an example of phenomenological kinetics, according to which the exponent of a power law represents the molecularity of a reaction, and accounts for the nonlinear dependence of fluxes on the concentration of intermediaries in a network (18). However, phenomenological models, by not considering a molecular mechanism specific to the reaction or the process being modeled, cannot be adequately tested and therefore have limited applicability. Our approach consists of modular building of models, in which each module represents the known or hypothesized kinetic scheme available for that process (16, 19–21). In the modular approach of computational modeling, a module is an edge linking at least two nodes, in analogy with the topological view of networks conceived as a collection of edges and nodes exhibiting large-scale

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organization (22, 23). Our modular approach, however, takes account of functional and regulatory interactions of reconstructed networks, a decisive feature for analyzing the dynamics of complex physiological responses. The modular approach of computational modeling also allows zooming in and out in a network of processes, this meaning that it is possible to choose how much detail the modeler will like to include. For instance, the tricarboxylic acid cycle (TCA cycle), or the glycolytic pathway may be described as set of reactions (8 or 10 reactions, respectively) or as a single aggregated step. Or a biochemical reaction may be written down considering the elementary kinetic steps of the catalytic cycle (24). New spatio-temporal organization properties appear when the modules are assembled into a unified scheme, that are considered emergent because they are unique, and thus could not had been anticipated from the properties of the isolated modules. In the following sections, we describe the modular approach followed to build models as applied to the bio-chemistry and -energetics of mitochondrial function. The following set of criteria specify the reliability of a computational model (1) sound physicobiochemical basis, (2) ability to reproduce qualitative and quantitative experimental data, (3) provision of meaningful explanation of the simulated experimental behavior, and (4) predictive power (4). Along the build-up and testing of our models we emphasize how we account for these criteria of reliability.

2. Materials 1. Basic biochemical information about the metabolic pathways and transport processes that are being modeled. This information can be obtained in biochemistry or physiology textbooks and some biochemical databases such as: ●

KEGG, Kyoto Encyclopedia of Genes and Genomes (http://www.genome.jp/kegg/).

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BRENDA, a comprehensive enzyme information database (http://www.brenda-enzymes.org).

2. Repository databases of models are a source for building our own model, constituting a good starting point for a beginner in the field of computational modeling. Many repositories of models are available, among them: ●

EMBL-EBI European Bioinformatics Institute (http:// www.ebi.ac.uk/biomodels-main/).

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The IUPS Physiome Proyect repository (http://models. cellml.org).

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ModelDB (http://senselab.med.yale.edu/ModelDB/).

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Pathways Logic Models (http://mcs.une.edu.au/~iop/ Bionet/index.html). The models in these databases are stored according to a series of standards in language, annotations (e.g., SMBL, CellML) and are publicly available for download.

3. Experimental data to constrain the modules’ kinetics, usually obtained under in vitro conditions (see below), and data to constrain the assembled model are available as well. 4. Computational tools to work with the modules, such as Matlab (www.mathworks.com), Mathematica (http://www.wolfran.com), Maple (http://www.maplesoft.com). These tools may be used in the modular analysis, while the model is built up, or with the assembled model. Some useful computational modeling packages have been developed for, e.g., Matlab. Among them the graphical package MatCont (http://www.matcont.ugent.be) is able to simulate time-dependent behavior and calculate the stability and the parameter sensitivity of the model (25).

3. Methods 1. Clear identification of the level of organization (e.g., molecular, (sub)cellular, (multi)cellular) being modeled, which should suit the question(s) addressed by the model. An important decision is whether the model will take into account spatial coordinates. This choice will determine if the model will be represented by a set of ordinary differential equations (ODEs, the only independent variable is time), or by partial differential equations (with time and spatial coordinates in one, two, or three dimensions, act as independent variables). We restrict the scope of this chapter to the discussion of models of ODEs (15). 2. Identification of the set of processes of interest along with observables (experimental variables) that have to be included in the model. For example, if the interest is to model the movement of a noncharged compound across the mitochondrial inner membrane, we need to account for the concentration gradient of the compound itself, and the kinetics of the transporter if mediated by a carrier, or the diffusion coefficient if moving through simple diffusion. However, for modeling the movement of a charged molecule or ion, the contribution of other charged compounds participating in the transport process, along with the driving force (mitochondrial membrane potential, pH gradient), will need to be considered as well. 3. Choice of the kinetic expressions describing the rate of the processes identified in point (2). According to our modular

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approach, the rate expression that best represents what is known about the mechanisms underlying the behavior of a particular step, will be chosen. Kinetic models of transport processes may encompass, for example, the Goldman–Hodgkin–Katz voltage equation (26, 27), or the Fick’s laws of diffusion for noncharged molecules following a concentration gradient in a homogeneous medium (28). Models will not always be mechanistically sound. Many processes have not been yet quantitatively and/or kinetically studied, to enable a mechanistic description. In such cases, an “ad hoc” equation may be written down, or a hypothetical functional relation. Savageau had proposed a power-law formalism postulating that each rate expression follows fractal kinetics where the order of the reaction with respect to each substrate/effector is the fractal dimension of the process (18). An attempt of a general formalism has been recently proposed, according to which kinetic models may be constrained using perturbation data (29). 4. Using your favorite program (MatLab, Maple, Mathematica, etc.), the first step is to represent the kinetic behavior of a particular module as a function of each one of the variables participating in the rate equation (16, 20, 26, 30, 31). Critical in this procedure is to choose the right set of parameters, which in the case of, e.g., an enzyme following Michaelis–Menten kinetics, correspond to the KM values for each of the substrates, products, or effectors participating in the equation, and the Vmax value of the enzyme at saturating levels of all substrates and/or effectors. When available, a first hint about parameter values can be obtained from experimental data (32). These values may need some adjustment since the conditions in which they have been measured may not correspond to physiological ones (see Subheading 4). As an example of a mechanistic rate equation, we consider the ATP hydrolysis associated with the activity of the actomyosin ATPase in cardiac muscle myofibrils (Fig. 1). The kinetics of this enzyme was described according to a Michaelis–Menten mechanism (30). The saturable dependence of the enzyme activity on ATP concentration, represented by the Lineweaver–Burk plot of the experimental data (Fig. 1a), and the competitive ADP inhibition (Fig. 1a, b), justify the rate expression adopted for this process (see ref. 20). 5. The sensitivity of the curve relating the rate of a given process, versus substrate or effector concentration to the values of the kinetic parameters, need to be studied. This will provide information about ways to modify a module behavior in the fully assembled model when the simulated output differs from experimental data. Studying the sensitivity of, e.g., the enzyme to parameters or variables also provides information about

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Fig. 1. Comparison of modeled and experimental behavior of myofibrillar ATPase.The figure shows (a) the Lineweaver–Burk representation of the experimentally measured rate of myofibrillar ATPase as a function of ATP concentration (36), whereas (b) shows the simulation of the experimental data after applying the equation on top of the figure. The competitive inhibition by ADP is shown by the addition of 0.5 or 2.0 mM ADP. The plots were produced with a maximal rate of the myofibrillar max ATPase, VAM , of 2.88 s−1, with P1, P2, and P3 representing the forms of tropomyosin that belong with an actomyosin complex with one, two or three cross-bridges, respectively. f01, f12, and f23, represent rate constants associated with the formation of cross-bridges. For parameter values and more detailed explanation see (20). (a) Is from Karatzaferi et al. (36) used with permission from the American Physiological Society.

mechanisms through which its activity might be modulated in vivo. As an illustration of this point, Fig. 2 shows the plots of the rate of isocitrate dehydrogenase as a function of isocitrate at two levels of Ca2+ (enzyme activator) and for different values of the activation constant for Ca2+, KCa. The decrease in KCa, allowed to simulate the stimulation of TCA cycle flux, within physiological Ca2+ levels in the mitochondrial matrix, and at different stimulation frequencies in ventricular cardiomyocytes. 6. The individual behavior of a module should be compared with experimental data obtained in vitro, from the isolated activity measured either in crude extracts, permeabilized cells, tissues, or partially purified. Eventually, in vivo experimental data can be obtained for some biological processes, and used to test the equation standing for the description of that process. The agreement of theoretical and experimental data may be qualitative, quantitative or, at best, both. 7. The modules may be assembled once their individual behavior is proved satisfactory according to the above criteria 1–6. At this stage, it may be convenient to schematize the relationship

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Fig. 2. Sensitivity of the plot of isocitrate dehydrogenase rate versus isocitrate to the level of Ca2+ and the Ca2+ activation constant. The activity of isocitrate dehydrogenase was studied as a function of the substrate, isocitrate, and at different levels of the activating Ca2+. To achieve a good reproduction of the stimulatory effect of Ca2+ as has been reported by (37), the Ca2+ association constant KCa (see equation on top of the plot ) was varied to 0.5 μM with respect to the original level reported in ref. 16.

among modules in a diagram such as shown in Fig. 3 for the model of mitochondrial energetic (16). The next step is to decide which of the variables participating in the equations will become state variables (i.e., whose values will inform the status of a system), and which will be adjustable parameters. For each state variables an ODE may be written with the following general scheme: dM = +Vproduction ± V transport - Vconsumption dt where M represents an intermediary (metabolite) in the system, or in a compartment thereof, Vproduction, the sum of the rates of all processes contributing M, Vconsumption, the sum of the rates of all processes consuming M, and Vtransport, the rates of the process that carry M inside or outside the compartment, or system being modeled. Table 1 shows the system of ODEs used in our model of mitochondrial energetics (16). Worth of notice at this stage is that the rate in ODEs may be subjected to scaling factors to account for volume or buffering effects. In Table 1, the buffering of the concentration of mitochondrial-free Ca2+ is taken into account by a constant factor, f [Table 1, (12)]. In the case that several compartments are

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Fig. 3. Schematic representation of the mitochondrial energetics model. Depicted are the mitochondrial electrophysiological and metabolic pathways, encompassing oxidative phosphorylation and matrix-based processes, along with their interactions, accounted for by the model. The tricarboxylic acid (TCA) cycle in the mitochondrial matrix is fed by acetyl CoA (AcCoA), which represents the point of convergence for fatty acids and glucose oxidation pathways, the two main fuels of the heart. The TCA cycle oxidizes AcCoA to CO2 producing NADH and FADH2 which provide the redox driving force for oxidative phosphorylation. NADH and FADH2 are oxidized by the respiratory chain and the concomitant pumping of protons across the mitochondrial inner membrane establishes an electrochemical gradient, or proton motive force (ΔμH), composed of electrical (ΔΨm) and proton (ΔpH) gradients. The ΔμH drives the phosphorylation of matrix ADP to ATP by the F1F0-ATPase (ATP synthase). The large ΔΨm of the inner membrane (−150 to −200 mV; matrix negative with respect to the cytoplasm) also governs the electrogenic transport of ions, including the cotransport of ATP and ADP by the adenine nucleotide translocator, Ca2+ influx via the Ca2+ uniporter and Ca2+ efflux via the Na+/Ca2+ exchanger (21). The model also accounts for the explicit dependence of the TCA cycle enzymes isocitrate dehydrogenase and α-ketoglutarate dehydrogenase on Ca2+. In this way, the rate of Ca2+ uptake by mitochondria is involved in membrane energization through the TCA cycle and oxidative phosphorylation. Key to symbols: ΔΨm is represented by the concentric circles with an arrow across located at the inner mitochondrial membrane. Reproduced from Cortassa et al. (16) with permission of Elsevier Science and Technology Journals in the format other book via Copyright Clearance Center.

considered, the volume ratio of the compartments has to be used to scale the rates if they belong to different compartments. 8. In order to perform a simulation, we need to consider the format with which to write the differential equations. The best advice there is to take a sample model from the chosen simulation package, and modify it to include the specific ODE from your own model. Then, parameter files have to be assembled and linked or included, as well as a set of initial values of the state variables (initial conditions). 9. Another important issue is the choice of the integration algorithm that will be used to simulate the model’s behavior. Among the matters to be considered for choosing the integration algorithm, include the “stiffness” of the model, i.e., if the time scale in

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Table 1 Mitochondrial energetic model ordinary differential equations State variable

Differential equation

[ADP]m

d[ADP]m V = ANT` VATPase` VSL dt dDY m VHe + VHe(F) - VHu - VANT - VHLeak - VNaCa - 2Vuni = dt Cmito

(1)

d[NADH] = -VO2 + VIDH + VKGDH + VMDH dt d[ISOC] = VACO - VIDH dt d[aKG] =VIDH VKGDH +VAAT ` dt

(3)

ΔΨm

[NADH] [ISOC] [αKG]

[ScoA]

d[SCoA] =VKGDH VSL ` dt

[Suc]

d[Suc] =VSL VSDH ` dt d[FUM] = VSDH - VFH dt d[MAL] = VFH - VMDH dt

[FUM] [MAL]

(2)

(4)

(5)

(6)

(7) (8) (9)

[OAA]

d[OAA] = VMDH - VCS - VAAT dt

(10)

[ASP]

d[ASP] = VAAT - VC _ ASP dt d[Ca 2 + ]m = f (Vuni - VNaCa ) dt

(11)

[Ca2+]m

(12)

For further details concerning the definition of each rate see ref. 16

which different processes attain steady state is very different (>102 of the time unit used), usually milliseconds for channel gating events and seconds for mitochondrial energy transduction processes. If some rapid kinetics is being accounted for, such as fast ion-transport mechanisms, then “stiff” solvers may be required. Most ODEs solver packages have their own integrators. Matlab includes a series of integrators for nonstiff ODEs systems (ode45, ode23, ode113), or for stiff ones (ode15s, ode23s), to name but a few. Fourth-order RungeKutta and Euler are commonly used integrators for nonstiff equations.

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10. Simulate the behavior of the assembled model as a function of time; first for a very short time (depending on the temporal resolution of the model may be a few ms, min, or hours) to see the stability of the model. To watch is whether some state variables take negative or very large or small values, or if they exhibit large deviations from physiological values. This step, which is the most experience-demanding one for the modeler, will provide key insights into which and how (extent and direction of change: increasing or decreasing) parameters may be adjusted to render reasonable simulations of the model behavior. 11. The next step is to run a model simulation till it reaches a steady state, i.e., where the derivatives of the state variables are less than 10−6–10−8 in relative terms (Δx/x). For the first simulations this may take a long time, depending on the initial values of the state variables. 12. At this stage, the modeler should start comparing the model output with experimental data (e.g., steady-state values of variables, or fluxes). Such comparison will lead to the finetuning of parameters, and the criteria utilized to do the parameter adjustment will need to be made explicit (see Note 2 on this subject). 13. When available, experimental data from transient behavior should be compared with model simulations, obtained under parametric conditions that reproduce those in which the experiment was carried out. As an illustrative example, Fig. 4 shows the temporal profile of bioenergetic variables (NADH, ΔYm) from isolated mitochondria. The similarity between the experimental and simulated behavior, qualitatively and quantitatively, indicates that the model accounts for the mechanisms operating in mitochondria when challenged with substrates, ADP, and uncouplers under in vitro conditions. The experimental–theoretical agreement achieved in the time-dependent behavior is of utmost importance since it is the performance of the model during transients that provides the most stringent test of its ability to represent physiological mechanisms. In the case of the ECME model (20), the dynamic response of mitochondrial energetics, following changes in workload conditions in heart trabeculae, was a main test of the model’s capacity to discriminate between competing hypotheses regarding the mechanisms underlying transient energy supply-demand matching. This is precisely what we mean by “explanatory power,” as mentioned in the introductory section. Utilizing the ECME model, the coupling between energy supply and demand, being hypothetically mediated by either

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Fig. 4. Time-dependent behavior of mitochondrial energetic variables during the state 4 → state 3 transition. Left panel shows the dynamic response of the endogenous fluorescent signal derived from NADH when a suspension of mitochondria were challenged by successive addition of glutamate plus malate (G/M, 5 mM each) or ADP (1 mM). Right panel shows the corresponding traces of ΔΨm as obtained from the calibrated ratiometric method with TMRM (tetramethylrhodamine methyl ester), and measured simultaneously with NADH. Experimental data are represented by filled squares, whereas the continuous (a) and dashed (b) lines stand for the simulated behavior obtained with the mitochondrial model of energy metabolism. Very recently, this model has been extended to comprise Na+, H+, Pi, and pH as state variables, including the regulation of mitochondrial activities by pH (40).

ATP or Ca2+, was tested during transients in which the temporarily increased workload imposed on the cardiomyocyte was simulated by means of a higher stimulation frequency (20, 33) (Fig. 5). Under those conditions, the involvement of ATP was investigated by modifying the rate constant of ATP hydrolysis by the myofibrils, which represents the most conspicuous source of ATP consumption in the working state. Similarly, in order to investigate the involvement of Ca2+, the maximal transport rate of Ca2+ through the mitochondrial Ca2+ uniporter was decreased. In each situation, simulations of the temporal profile of mitochondrial NADH and Ca2+ were obtained, and conclusions drawn (see caption of Fig. 5). 14. Regarding the “predictive power” of a model, it will be necessary to go beyond the traditional interpretation of the word “prediction.” Understandably, a model is able to predict if it can simulate a behavior that has not yet been observed. In the example shown in Fig. 6, the oscillatory behavior of reduced glutathione, GSH, was first observed in the model (Fig. 6a), and later confirmed experimentally (Fig. 6b) (31). Broadly speaking, if an integrated model is able to reproduce a behavior, whose mechanism was not intrinsically built in, even if such behavior was known beforehand, then it can be considered that the model is able to predict it.

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Fig. 5. Effects of modifying the activities of myofibrillar ATPase and the mitochondrial Ca2+ uniporter in the ECME model energetic behavior following changes in workload (20). (a) Shows the NADH behavior when the ECME model was studied during transitions in pacing frequency. Before region I, 0.25 Hz pacing was applied whereas in regions I and II the pacing frequency was kept at 2 Hz and decreased back to 0.25 Hz in region III. The black trace shows the effect of lowering the max maximal rate of ATP hydrolysis by the AM-ATPase to one half (VAM = 3.6 ´ 10-3 mM/ms) with respect to the control simulation (gray trace); all other parameters being identical in the two simulations. (b) Depicts the average profile of cytoplasmic ATP, ATPi, following changes in workload under control (gray trace) or decreased myofibrillar ATPase activity (black trace) according to the same protocol described in (a). (c) Exhibits the NADH profile obtained when the ECME model was studied with the same pacing protocol described in (a) but decreasing the maximal rate of the mitochondrial Ca2+ uni uniporter to one-tenth (black trace) (Vmax = 2.75 ´ 10-3 mM/ms) , with respect to the control (gray trace) simulation. (d) Shows the corresponding profile of mitochondrial Ca2+ when the uniporter activity was reduced in the simulation depicted in (c). Notice the strong reduction in [Ca2+]m accumulation (black trace) compared to the control simulation (gray trace). Reproduced from (20). with permission of Elsevier Science and Technology Journals in the format other book via Copyright Clearance Center.

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Fig. 6. Glutathione oscillations. (a) Displays the simulation of reduced glutathione (GSH) oscillations (100 s period) obtained under conditions of ΔΨm oscillations. (b) Shows the experimental demonstration of GSH oscillations (70-s period) recorded simultaneously with ΔΨm. Freshly isolated cardiomyocytes were loaded with 100 nM TMRM (ΔΨm probe) and 50 μM MCB (glutathione reporter) as described (31, 38). Oscillations were triggered after a localized laser flash, as previously described (39). Arrow indicates the timing of the flash. The decrease in the GSB signal corresponds to a drop in the GSH pool. Reproduced from (31) with permission of Elsevier Science and Technology Journals in the format other book via Copyright Clearance Center.

4. Notes 1. A crucial issue regarding the values of parameters and state variables concern the physical units used to represent them (19) see Chapter 2 there in. Modeling packages and numerical integrators work with numbers, disregarding their physical or physiological meaning. It is the modeler’s responsibility of providing equations with compatible units, which will render meaningful simulations. It is in modeler’s hands to interpret and give meaning to the output of a model. Experimental biologists, who sometimes do not trust quantitative computational models, often dismiss them with the expression “garbage in, garbage out”. Only a rigorous work with the layout and input of a model can disproof such claims. In our 20 years of work with various modeling approaches, we have seen many instances of units’ compatibility neglect that a simple troubleshooting could have exposed. 2. Regarding the issue of adjusting model parameters, the main inconvenient appears when different sources of data render very dissimilar values. Different sources, regarding organism or tissue from which the activity of interest has been isolated and/ or determined, or different assay conditions, can provide widely different results vis-â-vis, e.g., maximal rates a system is able to sustain. As a parameter, the Vmax of a process is the most likely candidate to be adjusted had an inconsistency in the dynamic behavior of the assembled model been detected. Table 2 shows data from (34) regarding experimentally measured differences in Vmax values of TCA cycle enzymes between in vitro and

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Table 2 Comparison of Vmax values of enzymes in vivo and in vitro from isolated enzyme kinetics determinations Enzyme

Vvivo (mM/min)

Isocitrate dehydrogenase α-Ketoglutarate dehydrogenase Malate dehydrogenase

Vmax (mM/min)

271.0

1.8

7,608.0

4.8

77.8

196

Malic enzyme

3.08

Succinate dehydrogenase

3.15

45

Citrate synthase

8.23

1,071

Pyruvate dehydrogenase

258.0

8.6

2.1

Data presented in ref. 34 where the Vvivo shows the in vivo values of maximal rates of enzyme obtained from NMR experiments with Dictyostelium discoideum TCA cycle dynamics, whereas Vmax stands for the purified enzyme maximal rate under in vitro conditions

in vivo determinations. It can be recognized that the differences may be up to three orders of magnitude as in the case of pyruvate, isocitrate, and α-ketoglutarate dehydrogenases as well as citrate synthase. These examples indicate that the in vitro condition provides an initial guess rather than an accurate figure of a parameter value. A more reliable estimation should be possible to obtain from parameter values which simulate more accurately experimental data with the computational model. 3. With the advent of Systems Biology, computational modeling is utilized to interpret and understand the physiological meaning of the information provided by high-throughput technologies such as genomics or proteomics. However, when multiple polypeptides are responsible of a single activity, such as in complex I in the electron transport chain, it is very difficult to predict what the effect on the activity will be when changes reported by proteomics in, e.g., polypeptide abundance of different magnitude and direction, are observed (see table 1 in (5)). In those cases, there are several criteria that should orient the course of action. First, it is necessary to know if the process in question exerts control over the fluxes in the network of interest (35). That means if the activity of the process is modulated, how much that will influence the fluxes throughout the network. This can be achieved with control analysis, in the biological system, by any of the methods developed to deal with the control of metabolic networks (35). In case the control is large, then, it will be necessary to estimate experimentally the impact of changing the subunit composition of the protein or complex on its activity.

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Another strategy is to test by model simulations which consequences bring about a parameter variation representing that activity. This is equivalent to applying control analysis to the model describing the metabolic network. If the effects of modulating the activity of interest are large, it would be justified to perform a detailed study regarding the influence of subunits composition on the activity of the enzyme or complex.

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31. Cortassa S, Aon MA, Winslow RL, O’Rourke B (2004) A mitochondrial oscillator dependent on reactive oxygen species. Biophys J 87:2060–2073 32. Barthelmes J, Ebeling C, Chang A, Schomburg I, Schomburg D (2007) BRENDA, AMENDA and FRENDA: the enzyme information system in 2007. Nucleic Acids Res 35:D511–D514 33. Brandes R, Bers DM (2002) Simultaneous measurements of mitochondrial NADH and Ca(2+) during increased work in intact rat heart trabeculae. Biophys J 83:587–604 34. Wright BE, Butler MH, Albe KR (1992) Systems analysis of the tricarboxylic acid cycle in Dictyostelium discoideum. I. The basis for model construction. J Biol Chem 267:3101–3105 35. Cortassa S, O’Rourke B, Winslow RL, Aon MA (2009) Control and regulation of mitochondrial energetics in an integrated model of cardiomyocyte function. Biophys J 96: 2466–2478 36. Karatzaferi C, Myburgh KH, Chinn MK, Franks-Skiba K, Cooke R (2003) Effect of an ADP analog on isometric force and ATPase activity of active muscle fibers. Am J Physiol 284:C816–C825 37. Rutter GA, Denton RM (1988) Regulation of NAD+-linked isocitrate dehydrogenase and 2-oxoglutarate dehydrogenase by Ca2+ ions within toluene-permeabilized rat heart mitochondria. Interactions with regulation by adenine nucleotides and NADH/NAD+ ratios. Biochem J 252:181–189 38. Aon MA, Cortassa S, Maack C, O’Rourke B (2007) Sequential opening of mitochondrial ion channels as a function of glutathione redox thiol status. J Biol Chem 282:21889–21900 39. Aon MA, Cortassa S, Marban E, O’Rourke B (2003) Synchronized whole cell oscillations in mitochondrial metabolism triggered by a local release of reactive oxygen species in cardiac myocytes. J Biol Chem 278:44735–44744 40. Wei AC, Aon MA, O’Rourke B, Winslow RL, Cortassa S. (2011) Mitochondrial energetics, pH regulation, and ion dynamics: a computational-experimental approach. Biophys J 100:2894–903

INDEX A Absorbance .................................... 21, 60, 63, 104, 105, 201, 209, 239, 245, 252–257 Acceptor ............................................. 7, 8, 90, 251, 252, 282 Acide crayeux ......................................................................2 Adenine nucleotide translocator (ANT) .............. 20, 29–30, 45, 59, 93, 98, 139, 208, 236, 237, 318 Adenosine 5’-diphosphate (ADP) .......................2, 9, 16, 18, 19, 20, 21, 27, 29, 31, 32, 33, 35, 37, 38, 40, 41, 43–46, 48, 49, 59, 61, 62, 64, 65, 66, 72, 91, 92, 95, 98, 103, 106, 110, 113, 114, 139, 140, 141, 148, 150, 162, 167, 171, 174, 175, 176, 189, 209, 243, 274, 282, 291, 292, 315, 316, 318, 319, 320, 321 ADP-activated respiration................................................. 59 ADP/O ................................................................. 15, 18, 19 ADP phosphorylation .................... 8, 18, 103, 104, 174, 181 ADP-regenerating systems .......................140, 141, 150, 176 Aerobic eukaryotic cells .......................................................7 α-ketoglutarate ................... 43, 106, 179, 288, 300, 318, 324 α-ketoglutarate dehydrogenase........................ 179, 318, 324 Alkylation .................................................260–261, 264, 265 Amplex Red.............................. 166, 167, 177, 178, 185, 189 Anode.. ............................................... 15, 16, 21, 22, 51, 170 ANT. See Adenine nucleotide translocator Antimycin A ................... 9, 20, 29, 31, 34, 35, 36, 42, 45, 91, 168, 178, 179, 185, 186, 188–192, 194, 203, 263, 266 Apoptotic cell death ........................................ 208, 209, 236 ARALAR ........................................................................ 219 Argon-krypton laser ........................................................ 245 Ascorbate........................................ 17, 19, 20, 29, 35, 42, 43 Aspartate (Asp) ................................. 42, 222, 274, 289, 290, 294, 295, 300, 305 ATP depletion ...................................................... 10, 208, 247 generation .................................................................. 238 synthase ................................. 4, 8, 16, 18, 19, 29–30, 44, 45, 89, 90, 91, 104, 119, 128, 133, 139, 148–149, 157, 162, 165, 171, 184, 189, 191, 208, 219, 243, 274, 282, 284, 318 synthasome ....................................................................4 synthesis .................................3, 4, 8, 33, 35, 89, 92, 123, 140, 162, 165, 171, 282, 283, 284, 287, 291, 292

ATPase ....................17, 59, 91, 137, 184, 315, 316, 318, 322 Atractyloside............................... 20, 30, 33, 93, 98, 209, 237 Average fluorescence intensity ......................................... 227 Avogadro constant ............................................................. 53 Azide... ......................................................................30, 34, 43

B Background ............................26, 39, 43, 47, 53–55, 79, 100, 136, 177, 178, 204, 223, 228, 229, 232, 245, 246, 247 Barometric pressure ..................................................... 28, 53 Basal respiration ...............................................60, 62, 64, 72 Bioactivity.......................................................................... 74 Biochemistry ................................................2, 250, 311, 313 Bioenergetics ..............................1–5, 9, 26, 47, 90, 136, 154, 188, 193, 201, 249–278, 281–306, 320 Biuret method ................................ 14, 15, 17, 156, 161, 216 Boltzmann’s constant ....................................................... 286 Bongkrekic acid ............................................................... 209 Brain mitochondria ................................11, 13–14, 185, 188 Brown adipose tissue ............................................... 153, 154

C Calcein fluorescence ........................................................ 238 Calcium ................................. 28, 93, 98, 137, 187, 197–199, 207–216, 219–241 Calcium green–5N .................................................. 239, 240 Calcium ionophore .......................................................... 227 Calcium overload..................................................... 141, 220 Calcium selective electrode ...................................... 210, 213 Calcium uniporter ....................................................... 93, 98 Calcium uptake......................... 199, 220, 223, 237, 240–241 Carbon......................................... 75, 76, 79, 80, 81, 86, 165, 166, 293, 294, 299, 302, 303, 304 Carbon fiber microelectrode ................. 74, 75, 79, 80, 81, 86 Carbonylcyanide p-trifluoromethoxyphenylhydrazone (FCCP).........................19, 20, 29, 31, 33, 44, 45, 67, 68, 69, 93, 95, 106, 107, 108, 109, 110, 111, 113, 114, 126, 127, 129–133, 156, 162, 167, 171, 173, 174, 175, 176, 181, 186, 188, 189, 192, 193, 194, 196, 200, 223, 253, 257 Carboxyatractyloside .................. 33, 106, 139, 141, 144, 146 Cardiac ischemia-reperfusion .......................................... 236

Carlos M. Palmeira and António J. Moreno (eds.), Mitochondrial Bioenergetics: Methods and Protocols, Methods in Molecular Biology, vol. 810, DOI 10.1007/978-1-61779-382-0, © Springer Science+Business Media, LLC 2012

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MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 328 Index Cardiac mitochondria ........................................................ 65 Cardiac muscle fibres....................................................... 315 Cardioprotective ...................................................... 236, 238 Catalase ..................................................27, 34, 51, 256, 258 Cation cycling ........................................................... 33, 139 Cationic fluorophores ...................................................... 244 Cationic indicators .......................................................... 124 CcO. See Cytochrome c oxidase cDNA.. ............................................................................ 156 Cell...... ........................... 2, 7, 26, 73, 89, 103, 119, 153, 166, 184, 207, 219, 236, 244, 249, 281, 316 Cell dysfunction .............................................................. 236 Cell membrane permeabilization..............27, 35–38, 41, 115 Cellular calcium homeostasis........................................... 207 Cellular respiration ...................................................... 73–87 Cell viability ....................................... 31, 33, 35, 36, 37, 163 Central nervous system...................................................... 74 Chelator........................................................................... 224 Chemical coupling hypothesis .............................................3 Chemical structures ......................................................... 281 Chemiosmotic hypothesis ...................................................3 5-(and–6)-chloromethyl–2’,7’-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA)............. 185 Citrate synthase activity..............................17, 21, 33, 34, 47 Citrate synthase ratio......................................................... 21 Citric acid cycle ........................................................... 3, 219 Clark-type oxygen electrode .......................18, 142, 186, 193 Clark-type polarographic oxygen sensors .......................... 51 CoASH ....................................................................... 97–99 Coenzyme Q ................................................... 154, 255–256 Complex I.................................4, 7, 8, 17, 19, 20, 42, 46, 90, 91, 171, 172, 179, 184, 213, 254, 255, 282, 324 Complex II .............. 4, 7, 17, 19, 20, 42, 46, 90, 91, 255, 282 Complex III ................................... 4, 8, 20, 90, 91, 178, 179, 181, 189, 255–256 Complex III (the cytochrome bc1 complex) ..................... 179 Complex IV .............................8, 17, 20, 90, 91, 95, 103, 256 Complex protein clusters .....................................................4 Concentration gradient ................................8, 139, 314, 315 Confocal imaging ............. 121, 226–227, 229–231, 245–247 Confocal microscopy ........................203, 219–234, 244, 247 Connectivity theorem ...................................................... 138 CoQ................................................................4, 42, 255, 256 CoQH2-cytochrome c oxidoreductase .................................4 Cozymase ............................................................................2 Crabtree effect ................................................................... 33 Creatine .............................140, 150, 151, 244, 289, 290, 305 Creatine phosphokinase .................................. 140, 141, 150 CyA..... .............................................................................. 99 Cyanide ................................................ 9, 34, 43, 91, 93, 233 Cyclophilin D .................................................................. 237 Cyclosporin A ...........100, 186, 193, 209, 210, 214, 215, 236 Cysteine residues ............................................................. 258 Cythochrome c-O2 oxidoreductase ......................................4

Cytoarchitectural organization .......................................... 74 Cytochrome aa3 content.................................................... 47 Cytochrome b..................................................8, 91, 253, 256 Cytochrome b560...................................................................7 Cytochrome c ............................... 4, 8, 17, 20, 27, 33, 37, 42, 43, 44, 47, 74, 90, 91, 208, 256–258, 276 Cytochrome c1 ................................................8, 91, 256, 257 Cytochrome c oxidase (CcO) .........................4, 8, 17, 33, 42, 74, 86, 90, 91 Cytochromes ......................................95, 252, 253, 256–258 Cytosol .................................................... 36, 37, 42, 91, 115, 186, 187, 191, 192, 197–199, 202, 219–225, 227, 228, 232, 236, 237, 238, 246, 266, 282, 284, 295

D Decoupled respiration ....................................................... 33 Dehydrogenases ........................... 42, 90, 138, 157, 179, 236, 250, 254, 255, 258, 274, 275, 282, 316, 317, 318, 324 Dephlogisticated air.............................................................2 Depolarization .........................................127, 128, 129, 131, 132, 199, 201, 208, 209, 220 Deuterium (2H) ............................................................... 281 Dichroic reflector............................................................. 245 Diffusion restriction .................................................... 44, 48 Digitonin ......................................... 9–11, 13, 19, 36–38, 40, 105, 106, 111–113, 115, 222, 259, 261, 262, 269 Digitonin-permeabilization ............................10–11, 13, 115 Dihydroethidium (hydroethidine) ................................... 186 Dim-ethyl sulfoxide (DMSO).......................65, 67, 70, 105, 106, 121, 155, 160, 167, 185–187, 221 Dissociation constant ...................................................... 220 DNP. See Dynamic nuclear polarization Donor... .................... 32, 41–43, 170, 171, 251, 252, 256, 283 Drug-induced toxicity ....................................................... 59 Dyes......................................... 104, 105, 106, 132, 203, 204, 209, 220, 221, 222, 223, 224, 226–228, 230, 231, 232, 233, 234, 240, 245, 268 Dynamic nuclear polarization (DNP) ..................... 295–298 Dyscoupling ................................................................ 33–35

E Elasticity coefficients ................................137, 138, 145–148 Electrical gradient ........................................................... 104 Electroactive species .................................................... 75, 79 Electrochemical backpressure ...................................... 34, 44 Electrochemical equilibrium.............................................. 92 Electrochemical gradient .................................3, 44, 89, 208, 219, 258, 282, 283, 318 Electrochemical potential ...............................3, 27, 119, 208 Electrochemical reduction ...................................................9 Electrochemical response ....................................................9 Electrometry ........................................................................5

MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 329 Index Electron ........................ 4, 7, 8, 15, 20, 29–30, 32, 33, 41–43, 45, 46, 47, 53, 55, 59, 74, 90, 91, 104, 119, 132, 133, 165–181, 189, 220, 250–256, 258, 259, 265, 275, 277, 282, 283, 284, 295, 296, 324 Electron donors ....................................41–43, 170, 171, 283 Electron leak............................................................ 165–181 Electron probe X-ray microanalysis................................. 220 Electron-transferring flavoprotein (ETF) .................. 29, 42, 44, 45, 46, 47, 275 Electron transfer system (ETS) ...............................8, 26, 27, 31–35, 44, 45, 46, 47, 55 Electron transfer system capacity ................................ 26, 32 Electron transport chain (ETC) ............................59, 62, 90, 119, 133, 175, 282, 283, 284, 324 Energy. .....................3, 4, 7, 26, 33, 89, 90, 91, 103, 104, 110, 114, 165, 174, 207, 243, 249, 252, 254, 258, 281, 282, 284, 285, 286, 295, 296, 312, 319, 320, 321 Energy metabolism.................................................. 258, 321 ETC. See Electron transport chain ETF. See Electron-transferring flavoprotein ETS. See Electron transfer system Excitation spectra .................................................... 229, 231 Extramitochondrial concentration ................... 105, 215, 235

F FAD 7, 103, 104, 266 Fatty acid β-oxidation ....................................................... 42 Fatty acid substrates........................................................... 42 FCCP. See Carbonylcyanide p-trifluoromethoxyphenylhydrazone FCR. See Flux control ratios Filter.......................................40, 62, 63, 66, 77, 83, 121, 124, 130, 131, 155, 160, 169, 194, 195, 197, 226, 228, 231, 245, 260, 263, 268, 277 Flavoprotein ....................................... 42, 253, 255, 256, 275 Fluo–4/AM ..............................................124, 221, 223, 225 Fluorescence ........................................ 10, 61, 105, 119, 177, 188, 219, 238, 245, 254 Fluorescent emission spectra ........................................... 245 Fluorescent/luminescent probes ..........................................7 Fluorescent probes ................................... 103–115, 219–234 Fluorophore ..............................................229, 244, 245, 269 Flux control coefficients ...................137, 138, 147–149, 151 Flux control ratios (FCR) .....................26, 32, 33, 34, 46–48 Flux control summation theorem .................................... 138 FMN proteins .....................................................................7 Free fatty acids ................................................11, 27, 93, 153

G Glucose (Glc) .........................2, 10, 12, 33, 35, 77, 103, 105, 121, 140, 150, 155, 166, 184, 192, 193, 194, 195, 197, 198, 200, 222, 244, 263, 266, 278, 282, 283, 289, 290, 298, 299, 300, 303, 305, 306, 318

Glutamate (Glu) .................................. 18, 19, 20, 25, 29, 37, 41, 42, 44, 45, 61, 62, 64, 65, 66, 72, 74, 93, 99, 106, 127, 128, 130, 131, 132, 150, 168, 179, 185, 189, 239, 259, 275, 288, 290, 293, 294, 295, 297, 300, 301, 303, 304, 305, 321 Glutamate/malate ......................18, 61, 62, 64, 65, 66, 72, 93 Glutamate receptor ............................................ 74, 128, 132 Glutamine (Gln) .....................................155, 184, 260, 263, 266, 278, 289, 290, 300, 305 Glutathione ..................................................... 261, 321, 323 Glycerophosphocholine (GPC) ............................... 289, 290 Glycine ......................127, 128, 130, 131, 133, 155, 259, 260 Glycolysis .................................................................... 3, 248 Glycolytic ATP........................................................ 133, 208 GPC. See Glycerophosphocholine GSH/GSSG .............................................251, 259, 261–263 Guanylate cyclase ........................................................ 73, 74

H Heart mitochondria .......................................11, 14, 17, 114, 139–144, 146, 150, 151, 207, 240 Helium-neon laser ........................................................... 245 Heme......................................................................... 34, 256 High energy compounds .....................................................3 High-resolution respirometry .............................. 25–55, 150 High-throughput analysis ........................................... 59–72 High-throughput method ................................................. 59 Hippocampal slice ............................................... 73–87, 299 Hippocampus ....................... 74, 83, 288, 289, 290, 304, 305 Histidine............................................................................ 43 HL–1 cells ............................................................... 223, 225 H2O2. See Hydrogen peroxide HPLC.. .............................. 254, 255, 259, 261–263, 272, 278 Human embryonic kidney cells (HEK293) ....... 32, 153–163 Hydrogen peroxide (H2O2)............................... 46, 185–188, 195, 200, 202, 264 Hyperpolarization ....................................127, 133, 198, 296 Hypoxia ................................................................... 246–248

I ICAT. See Isotope-coded affinity tag Immunoassay ................................................................... 250 Infrared range .......................................................... 229, 230 Inhibitory effect ......................................................... 69, 297 Inner mitochondrial membrane........... 27, 33, 41, 42, 89, 90, 91, 103, 104, 106, 109, 123, 198, 201, 208, 236, 318 Inorganic phosphate ...............................28, 43–45, 282, 291 Inositol (Ins) .................................................... 289, 290, 305 Intermediary metabolism......................................... 293, 306 Intracellular ................................... 7, 9, 45, 48, 73, 220, 222, 227, 238, 245, 247, 292 Intramembrane space ..........................................................8 Intrinsic uncoupling .................................................... 33, 34 Ion gradients...................................................................... 26

MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 330 Index Ionomycin........................................................................ 232 Ionophores.................................. 93, 100, 107, 150, 224, 227 Iron-sulfur proteins ............................................7, 8, 91, 275 Ischemia-associated pathologies ............................. 141, 209, 236, 237, 238, 297 Isotope-coded affinity tag (ICAT) .................. 261, 269–276 Isotopomer ....................................... 284, 294, 295, 303, 304

J JC–1 (5,5’,6,6’-tetrachloro–1,1,’3,3’tetraethylbenzimidazolylcarbocyanine iodide) Malate.................................................................. 185

K Krebs cycle.............. 3, 20, 103, 104, 138, 282, 283, 284, 288, 293–296, 302, 304, 306

L Lambda scan ................................................................... 228 Larmor frequency .................................................... 286, 296 Leak control ratio ........................................................ 34, 47 Leak state .........................................................31, 33, 45, 48 Leica confocal.......................................................... 226, 233 Linear regression ................................ 55, 67, 68, 69, 71, 213 Liver mitochondria .............................. 12, 14–15, 61–63, 65, 72, 94–95, 105, 114, 150, 151, 210, 215, 220 Living cells ...................................................................... 2, 5

M Magnetic field ..................................284–287, 295, 296, 304 Magnetic resonance spectroscopy (MRS)........ 283–289, 298 Magnets..............................................31, 284–286, 291, 304 Malate.................................18, 19, 20, 29, 37, 41, 42, 44, 45, 48, 49, 61, 62, 64, 65, 66, 72, 93, 99, 106, 141, 142, 149, 150, 167, 168, 172, 179, 181, 185, 189, 239, 275, 294, 295, 321, 324 Malonate ............. 20, 140, 151, 156, 157, 162, 167, 175, 176 Malonyl-Coa ..................................................................... 97 Mass-specific oxygen flux ...........................33, 40, 41, 46, 47 Mass spectrometry............................250, 254, 261, 269–273 Matrix space ...................................................................... 41 MCA. See Metabolic control analysis Membrane permeability subsystem ................................. 139 Membrane potential ............................26, 89–100, 103–115, 119–133, 139, 142–148, 150, 157, 158, 162, 167, 172–174, 183–204, 209, 213, 238, 243, 314 Metabolic control analysis (MCA) .................. 135–138, 141 Metabolic pathways .......................................2, 26, 288, 289, 294, 306, 313, 318 Metabolic rates ............................................................ 2, 284 Metabolic states .................................. 26, 41, 48, 53, 89–100 Metabolism ................................... 28, 86, 91, 221, 253, 258, 281–298, 306, 321 MiR06.. .........................27, 36, 39, 40, 41, 43, 44, 46, 49–54

Mitochondria ...................................... 1–5, 7–22, 26, 59–72, 74, 89–100, 103–115, 119–133, 135–151, 153, 165–181, 183–204, 207–216, 219–241, 243–278, 281–306, 311–325 Mitochondrial and cellular bioenergetics......................... 136 Mitochondrial bioenergetics ............................................ 1–5 Mitochondrial calcium accumulation .............................. 208 Mitochondrial calcium transport measurements ..... 213–215 Mitochondrial calcium uptake ................................. 220, 223 Mitochondrial complexes .................................................. 86 Mitochondrial complex III ..................................................4 Mitochondrial dehydrogenases ........................................ 236 Mitochondrial depolarization ...................129, 199, 208, 209 Mitochondrial dysfunction ................................................ 59 Mitochondrial electron transfer .............................. 165, 171, 174, 250, 252–254 Mitochondrial function ...............................38, 59, 249, 250, 251, 277, 311–325 Mitochondrial inner membrane ......................27, 33, 42, 89, 90, 91, 104, 109, 123, 139, 153, 158, 165, 171, 173, 174, 177, 179, 181, 201, 208, 314, 318 Mitochondrial intermembrane space .....................3, 90, 104, 157, 258, 282, 283 Mitochondrial isolation ........................................ 13–15, 36, 60–63, 69–72, 105, 107, 155, 161, 166, 168–169, 209–212, 269 Mitochondrial matrix ....................................8, 90, 123, 125, 126, 127, 173, 177, 186, 191, 219, 237, 318 Mitochondrial membrane potential.................... 26, 89–100, 103–115, 119–133, 139, 143, 144, 150, 172–174, 183–204, 238, 243, 314 Mitochondrial oxidative phosphorylation ....................... 220 Mitochondrial permeability transition pore (MPTP).........184, 208, 209, 213–215, 220, 235–241 Mitochondrial respiration................................16, 18, 19, 26, 27, 35, 41, 47, 52, 74, 86, 89, 114, 140, 144, 145, 146, 147, 148, 149, 150, 161, 162, 174, 248, 255, 258 Mitochondrial respiratory chain ......... 7, 8, 74, 200–202, 250 Mitochondrial ryanodine receptor ................................... 207 Mitochondrial swelling............. 109, 110, 114, 115, 236, 239 Mitochondrial uncoupling proteins (UCPs)............................................90, 153–163, 166 Mitochondria Na+-Ca2+ exchanger ...........235, 237, 238, 318 Mitochondrion .........................................7, 8, 207, 227, 277 MitoSox Red .............186, 187, 190–191, 197, 198, 202, 204 Mitotracker red ............................................................... 233 Modular kinetic analysis .................................................. 137 Molecular manipulations ................................................. 250 Molecular proton pumps .....................................................4 Molecular redox states ....................................................... 26 MPTP. See Mitochondrial permeability transition pore mRNA ............................................................................. 156 MRS. See Magnetic resonance spectroscopy Multiphasic oxygen kinetics .............................................. 49 Multiphoton confocal imaging ................................ 229–231

MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 331 Index Myocytes ..................................................237, 238, 244–247 Myxothiazol ..................................... 20, 30, 42, 45, 128, 140

N NADH .......................................3, 4, 5, 7, 20, 36, 42, 90, 91, 92, 104, 162, 179, 250, 253–256, 258, 259, 275, 277, 282, 283, 291, 318, 319, 320, 321, 322 NADH-CoQ oxidoreductase ..............................................4 NADH dehydrogenase .............................250, 258, 275, 282 NAD+/NADH.......................................5, 92, 179, 254, 259 NADPH..................4, 10, 253, 259, 261, 265, 277, 278, 291 NADP+/NADPH ............................................... 5, 258–259 Nafion®, 75, 76, 80, 81, 86, 87 Nernst equation ................................... 92, 94, 108, 112, 115, 173, 213, 252, 254, 255, 273 Nernst equilibrium .......................................................... 124 N-ethylmaleimide (NEM) ......... 99–100, 209, 260, 264, 265 Neuro modulator ............................................................... 74 Neuron .....................................................120, 125, 127–132 Nigericin........................................ 93, 95–96, 150, 156, 162, 167, 172, 173, 175, 176, 247 Nitric oxide (NO) ...........................................73–87, 91, 137 NMDA receptor ...................................................... 128–133 N,N,N’,N’-tetramethyl-p-phenylenediamine (TMPD) ............................. 17, 19, 20, 29, 35, 42, 43 NO microsensor ........................................76, 77, 79–82, 86 Non-descanned detector.................................................. 229 Normoxia........................................................................... 48 Nuclear magnetic resonance (NMR) spectroscopy ..................................254, 281–306, 324 Nuclear spins ............................................285–287, 295, 296 Nuclei.. ......................................... 94, 95, 211, 284, 288, 291

O Octanoylcarnitine .......................................29, 41, 42, 44, 45 O2/electron ratio ................................................................ 53 Oligomycin .................................9, 19, 20, 30, 31, 33, 34, 92, 96, 99, 106, 110, 123, 127, 128, 129, 130, 131, 133, 139, 140, 141, 144, 145, 147, 149, 156, 157, 162, 167, 171, 174, 175, 176, 181, 185, 186, 189, 190, 191, 192, 193, 194, 196, 197, 198, 203 o-phenylenediamine........................................................... 75 Organelle fluorescence..................................................... 228 OROBOROS Titration .................................................... 54 Oversaturation ................................................................. 245 Oxidative phosphorylation (OXPHOS)..................8, 19, 20, 25–55, 89, 112, 113, 135–151, 165, 168, 171, 172, 174–177, 202, 207, 219, 220, 249, 318 Oxidative processes..............................................................5 Oxidative stress.........................................113, 208, 209, 258 Oxidized ............................................................7, 36, 43, 91, 253, 258, 261, 263, 264, 265, 266, 269, 273–277, 282, 283, 318

OXPHOS. See Oxidative phosphorylation Oxygen concentration ................................. 18, 28, 31, 41, 43, 44, 46, 48–55, 60, 71, 143, 151, 180, 192, 193 consumption ....................................9–15, 17–21, 28, 31, 34, 43, 46, 53, 55, 59–72, 143, 144, 149, 151, 156–158, 162, 167, 170, 171, 175, 176, 181, 186, 192, 193, 257–258, 304 consumption measurements....................................... 192 conversion .............................................................. 68, 71 electrodes .................. 10, 11, 15, 16, 19, 21, 22, 142, 149, 170, 171, 172, 175, 176, 180, 181, 186, 193, 212 solubility .................................................... 21, 22, 52–54 superoxide .................................................... 91, 165, 177 Oxygen back-diffusion ................................................ 53, 54 Oxygine ...............................................................................2 Oxygraph chamber ...................................................... 28, 50

P Palmitic acid ...................................................................... 42 Palmitoylcarnitine.............................................................. 42 Pathophysiological uncoupling .......................................... 33 Perchloric acid (PCA) .....................................259, 262, 289, 290, 299–300, 302, 304 Perfused heart .................................................. 297, 304, 305 Permeability transition .................................... 113, 114, 208 Permeability transition pore ............................. 99–100, 184, 193, 208, 220, 235–241 Permeabilization medium .................................................. 10 Permeabilized cells ............................................9, 10, 19, 27, 35–51, 55, 103–115, 316 Peroxiredoxins (Prx) ......................... 264, 265, 268, 273, 277 pH....... ....................................................................3, 5, 10, 11 Pharmacological studies .................................................... 86 Phenomenological kinetic analysis .......................... 135–151 Phenylarsine oxide ........................................................... 209 pH gradient ............................................. 3, 91, 96, 104, 162, 172, 179, 208, 243, 314, 318 Phosphate .............................................. 3, 10, 28, 43–45, 54, 91, 95, 105, 114, 139, 140, 151, 208, 209, 237, 243, 260, 282, 291, 293, 300 Phosphate carrier ............................................. 139, 208, 237 Phosphocholine (PCho) .......................................... 289, 290 Phosphocreatine (PCre) ............... 10, 27, 284, 289–291, 305 Phosphorescent water-soluble oxygen probe ..................... 60 Phosphorus ...................................................................... 291 Phosphorylating subsystem ...... 139, 140, 141, 142, 144–148 Photobleaching........................................................ 229, 269 Pixel...... .............................223, 229, 231, 233, 245, 246, 247 Plasma membrane ..................................... 33, 35, 36, 37, 43, 104, 111, 120, 123–126, 128, 129, 131, 132, 223, 224, 227–228, 238 Plasma membrane permeabilization ....................36, 37, 223, 224, 227–228

MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 332 Index Plasma membrane potential indicator (PMPI) (anionic indicator) ............................... 120, 124, 125 Platinum cathode .........................................16, 22, 170, 180 Platinum electrodes .............................. 9, 15, 22, 75, 78, 170 Polarography ...................................................................... 60 Polymer films..................................................................... 75 Postconditioning.............................................................. 238 Preconditioning ....................................................... 137, 238 Probe... .................................11, 60, 61, 63, 64, 67, 68, 70, 71, 75, 103–115, 120, 121, 123, 124, 125, 126, 129, 130, 132, 168, 173, 177, 185, 189, 203, 216, 219–234, 239, 255, 261, 286–288, 306, 323 Prosthetic groups .................................................................7 Protein.. ..................................4, 7, 33, 60, 90, 106, 142, 153, 166, 187, 207, 235, 250, 282, 311 Proteoliposomes ...................................................... 105, 154 Proton circuit ............................................................. 33, 44, 119 concentration gradient ............................................... 139 electrochemical potential ............................. 33, 119, 208 imbalance .......................................................................8 leak ................... 33, 41, 44, 119, 133, 139, 140, 141, 142, 144–148, 153, 156–158, 161–162, 165–181, 252 leak kinetics .........156, 157, 158, 161–162, 168, 175–176 pumps ............................3, 4, 16, 17, 34, 41–43, 255, 256 Protonmotive force .............................. 3, 4, 16, 91, 103, 119, 136, 138, 139, 153, 173, 174, 175, 179, 243 Protonophore........................ 20, 33, 107, 111, 123, 129, 171 Pyruvate....................................... 19, 20, 29, 42, 43, 48, 106, 141, 142, 149, 155, 167, 172, 181, 222, 251, 275, 282, 294, 295, 297, 298, 300, 303, 305, 324

Q Q-junction ......................................................................... 42 Quantitative analysis linearise ........................................... 67 Quench ..................................... 123–126, 128–133, 228, 238 Quenching ................................................105, 106, 123, 178

R Radical species .....................................................................5 Radio frequency (RF) .......................193, 286–288, 301–303 Range indicator ............................................................... 229 “Rapid mode” RAM ........................................................ 207 Ratiometric imaging ........................................................ 244 RCR. See Respiratory control ratio Reaction medium ....................................10, 13, 93, 95–100, 106, 110, 111, 113, 114 Reactive oxygen species (ROS)........................33, 34, 86, 91, 166, 183–204, 238, 249, 252, 256 Red fluorescent dye ......................................................... 226 Redox balances ....................................................................5 Redox potentials .....................................5, 92, 250–254, 259 Redox proteomics .....................................258, 261, 269–273 Redox systems ..........................................3, 4, 250, 258, 278

Redox western blot ................... 250, 260–261, 263–267, 278 Reduced ................................15, 42, 49, 51, 54, 91, 103, 104, 163, 179, 189, 238, 253, 256, 257, 261, 263, 265, 266, 269, 273–278, 283, 287, 293, 300, 321–323 Reducing agents Reducing equivalents ...................... 4, 91, 165, 255, 282, 284 Region of interest (ROI) ...........................84, 130, 131, 223, 227, 228, 229, 232, 286, 287, 288 Reoxygenation ............................................43, 44, 48, 49, 51 Resazurin ......................................................... 187, 200, 202 Resolution ....................................... 4, 19, 25–55, 70, 75, 84, 86, 121, 123, 124, 130, 131, 150, 232, 237, 289, 293, 297–302, 304, 306, 320 Respiratory buffer .......................................19, 61, 63, 64, 70 Respiratory chain............................... 7, 8, 16, 18, 19, 20, 74, 90, 91, 93, 104, 128, 138, 140, 144, 147, 175, 184, 187, 189, 200–202, 208, 250, 252, 258, 277, 318 Respiratory complexes ......................... 21, 27, 104, 165, 168, 171, 172, 177, 179, 213 Respiratory complex I......................................104, 165, 168, 171, 172, 177, 213 Respiratory Complex I (NADH:ubiquinone oxidoreductase) .................................................... 179 Respiratory control ratio (RCR) ........................... 15–18, 34, 48, 62, 69, 72, 156, 161–163, 171, 176 Respiratory flow ................................................................ 33 Respiratory flux control ratios ........................................... 47 Respiratory subsystem .............. 138, 139, 142, 144–148, 151 Respirometric viability index ....................................... 35–36 Respirometry ............................................................... 25–55 RF. See Radio frequency Rhod–2 ............................................................ 220–232, 234 Rhodamine 123 (R123) ............................104, 121–132, 244 ROI. See Region of interest ROS. See Reactive oxygen species Rotenone ................................. 20, 29, 31, 34–38, 43–45, 67, 91, 93, 95–99, 140, 141, 144–147, 150, 156, 162, 167, 168, 171, 172, 175, 176, 179, 181, 210, 213, 215, 263 Routine respiration .......................................... 26, 31–35, 37 Ruthenium red ............................................................ 93, 98

S Safranine ..........................................103–115, 185, 189, 190 Salicylate/Br-Succinimide ..................................... 93, 96–97 Saponin .................................... 9–13, 19, 36, 39, 40, 41, 222 Saponin-permeabilization ............................... 10, 12–13, 40 Screening ................................ 60, 61, 62, 64–67, 69, 72, 209 Selective membrane permeabilization .................................9 Single-to-noise ratio ........................................................ 245 Skeletal muscle fibres................................................... 36, 55 Skeletal muscle mitochondria ......................61, 63, 154, 166, 168, 171, 172, 173, 174, 175, 176 SNARF–1 ............................................................... 243–248

MITOCHONDRIAL BIOENERGETICS: METHODS AND PROTOCOLS 333 Index Sodium dithionite (Na2S2O4) ..............................52, 54, 167, 170, 180, 253, 257 Spectroscopy ..................... 254, 257, 284–289, 291, 304–305 Spectrum ................................. 167, 221, 289, 290, 291, 292, 295, 299, 300, 302, 303, 305 Sprague-Dawley rats ......................................................... 70 Stable isotopes ......................................................... 296, 298 Standard curve ................................................... 63, 178, 247 Standard polarography ...................................................... 60 State 2........................................16, 18, 62, 64, 65, 67, 68, 69 State 3.................................................. 16–18, 33, 45, 48, 62, 64–69, 139, 140, 142, 147, 148, 149, 171, 174, 175, 181, 321 State FCCP ....................................................................... 19 State oligomycin ................................................................ 19 State 3 respiration..................16, 17, 18, 62, 64, 68, 140, 149 Steady-state ................................... 22, 85, 97–100, 107, 108, 110, 111, 113, 126, 137, 138, 140, 142, 143, 148, 157, 162, 174, 175, 181, 207, 236, 237, 250–257, 259, 269, 277, 278, 292, 299, 303, 312, 319, 320 Steady-state redox potentials ........................... 250–254, 259 Substrate oxidation ............................................8, 10, 15, 35, 103, 142, 154, 157, 179 Substrate saturation ........................................................... 42 Substrate–uncoupler–inhibitor titration (SUIT) protocols .........................................29–30, 37, 44–47 Succinate ................................4, 7, 18, 20, 29, 31, 32, 35–38, 41–45, 49, 61, 62, 64–66, 72, 90, 93, 95, 98, 99, 140, 141, 150, 151, 156, 157, 162, 167, 168, 171, 173, 175, 176, 178, 179, 181, 185, 190, 210, 214, 215, 253, 275, 282, 324 Succinate-CoQ oxidoreductase ...........................................4 Succinate dehydrogenase .............. 42, 90, 157, 275, 282, 324 Superoxide ................................. 91, 165–168, 177–179, 181, 185, 186, 187, 189–192, 195, 197, 198, 202, 249 Superoxide dismutase ...................................... 167, 177, 181 Swelling ............. 109, 110, 114, 115, 208, 215, 236, 239, 247 Synaptosomes ............................................................ 74, 188

Texas red.......................................................................... 233 Thiol/disulfide state ........................................................ 261 Thiols.. ....................................................... 252, 266, 273, 277 Thioredoxin–2 (Trx2) ..............................250, 252, 260, 261, 263, 267–269, 273, 277, 278 Thioredoxin reductase (TrxR) ................................ 260, 261, 264–269, 277, 278 Threshold time .................................................................. 71 Tissue biopsies......................................................... 289, 298 Tissue extracts .........................................283, 284, 289, 291, 298, 299, 300, 301, 303, 304 Titanium stoppers ............................................................. 54 TMPD. See N,N,N’,N’-tetramethyl-p-phenylenediamine ‘Top-down’ modular kinetic (elasticity) analysis ...... 136–140 Total fluorescence ............................................................ 228 TPP+-selective electrode .......................................... 100, 142 Transmembrane electric potential ................................... 3, 8 Tricarboxylic acid cycle .................................36, 90, 313, 318 Tuneable laser .................................................................. 229

T

Wavelengths ............................ 105, 106, 109, 110, 131, 167, 177, 189, 190, 191, 192, 200, 204, 221, 222, 228–233, 245, 246, 247, 252, 253, 254, 257

Taurine (Tau)................................ 10, 27, 244, 289, 290, 305 TCA cycle ......................................... 36, 42, 93, 96, 97, 282, 283, 284, 297, 313, 316, 318, 323, 324 Terminal electron acceptor ..................................................7 Tetramethylrhodamine methyl ester (TMRM) ..............121–132, 186, 187, 192, 193, 194, 196–201, 203, 228, 244, 321, 323 Tetraphenylphosphonium (TPP+).................... 92–100, 109, 141, 142, 143, 144, 150, 209, 215

U Ubiquinol ..................................... 90, 91, 179, 255, 256, 276 Ubiquinone................................. 4, 42, 90, 91, 179, 255, 275 Ubiquinone pool...................................................... 3, 4, 179 Ubisemiquinone ...................................................... 255, 256 Uncoupled respiration ....................................................... 19 Uncoupling effect ........................................................ 42, 69 Undersaturation ............................................................... 245

V Valinomycin .............................................105, 106, 107, 108, 111, 112, 141, 143, 150, 247 Vasodilation ....................................................................... 74 VDAC. See Voltage-dependent anion channel Vector............................................................... 156, 286, 287 Vital principle ......................................................................2 Voltage-dependent anion channel (VDAC) ............ 237, 276

W

X Xenobiotics ........................................................................ 17

Z Zeiss 510 META confocal .............................................. 228 Zymase ................................................................................2