Mitochondrial Regulation: Methods and Protocols (Methods in Molecular Biology, 2310) 1071614320, 9781071614327
This fully updated edition explores the different pathways that converge into the regulation of mitochondrial function.
122 23
English Pages 330 [319] Year 2021
Preface
Contents
Contributors
Chapter 1: Isolation of Mitochondria from Liver and Extraction of Total RNA and Protein: Analyses of miRNA and Protein Express...
1 Introduction
2 Materials
2.1 Mitochondrial Isolation: Materials and Reagents
2.2 Total RNA Isolation: Materials and Reagents
2.3 miRNA Expression Analysis: Materials and Reagents
2.4 Total Protein Isolation: Materials and Reagents
3 Methods
3.1 Mitochondria Isolation
3.2 Total RNA Extraction from Isolated Mitochondria
3.3 miRNA Expression Analysis
3.4 Total Protein Extraction from TRIzol-Treated Isolated Mitochondria
4 Notes
References
Chapter 2: Determination of Oxidative Phosphorylation Complexes Activities
1 Introduction
2 Materials
2.1 Reagents and Buffers
2.2 Equipment Required
3 Methods
3.1 Isolation of Hepatic Mitochondria
3.2 Mitochondrial Quantification
3.3 Citrate Synthase Activity Assay
3.4 Individual Respiratory Chain Enzymatic Activities
3.4.1 Complex I (NADH:Ubiquinone Oxidoreductase)
3.4.2 Complex II (Succinate:Ubiquinone Oxidoreductase)
3.4.3 Complex III (Coenzyme Q: Cytochrome c Oxidoreductase)
3.4.4 Complex IV (Cytochrome c Reductase)
3.4.5 ATPase Activity of Mitochondrial ATPSynthase
4 Notes
References
Chapter 3: BN-PAGE-Based Approach to Study Thyroid Hormones and Mitochondrial Function
1 Introduction
2 Materials
2.1 Materials and Equipment
2.2 Mitochondria Isolation and BN-PAGE Sample Preparation
2.3 Gel Casting and Electrophoresis
2.4 Brilliant Blue G Gel Staining
2.5 Histochemical Evaluation of OXPHOS in-Gel Activities
3 Methods
3.1 Isolation of Mitochondria
3.2 Solubilization of OXPHOS Respiratory Complexes from Crude Mitochondrial Pellet
3.3 Casting of Acrylamide Gradient Gels
3.4 Electrophoresis
3.5 Gel Staining with Brilliant Blue G
3.6 Quantification of Native OXPHOS Protein in-Gel Activity by Histochemical Evaluation
4 Notes
References
Chapter 4: Monitoring Mitochondrial Function in Mouse Embryonic Stem Cells (mESCs)
1 Introduction
2 Materials
2.1 mESC Culture
2.2 Seahorse XFe24 Live-Cell Metabolic Assay
2.3 Flow Cytometry Analysis of Mitochondrial Transmembrane Potential
3 Methods
3.1 Seahorse XFe24 Live-Cell Metabolic Assay
3.1.1 Assay Preparation (the Day before) (See Notes 1 and 2)
3.1.2 Seahorse Assay Protocol
3.1.3 Normalization of the Seahorse Assay by Cell Count (See Note 6)
3.2 Flow Cytometry Analysis of Mitochondrial Transmembrane Potential Using the TMRM Probe
4 Notes
References
Chapter 5: Mitochondrial Functional Assessment in Mammalian Gametes Using Fluorescent Probes
1 Introduction
2 Materials
2.1 Cell Media
2.2 Fluorescent Probes for ROS Detection
2.3 Fluorescent Probes for MMP Assessment
2.4 Fluorescent Probes for Mitochondrial Mass/Distribution/Aggregation Assessment
2.5 Controls
2.6 Equipment
3 Methods
3.1 Sperm Cells
3.1.1 Sample Preparation
3.1.2 ROS Content Assessment
3.1.3 MMP Evaluation
3.2 Oocytes
3.2.1 Oocytes Preparation
3.2.2 Mitochondrial H2O2 Levels
3.2.3 MMP Evaluation
3.2.4 Mitochondrial Mass/Distribution/Aggregation
4 Notes
References
Chapter 6: Update on the Histoenzymatic Methods for Visualization of the Activity of Individual Mitochondrial Respiratory Chai...
1 Introduction
2 Materials
2.1 Assessment of Cytochrome c Oxidase (COX; Complex IV) Activity in the Human Frozen Sections
2.1.1 Solutions and Chemicals
2.1.2 Equipment and Accessories
2.2 Assessment of Succinate Dehydrogenase (SDH; Complex II) Activity in the Human Frozen Sections
2.2.1 Solutions and Chemicals
2.2.2 Equipment and Accessories
2.3 Assessment of NADH Dehydrogenase (Diaphorase; Complex I) Activity in the Human Frozen Sections
2.3.1 Solutions and Chemicals
2.3.2 Equipment and Accessories
3 Methods
3.1 Brain, Heart, Liver Autopsy Samples and Muscle Biopsy from Patients
3.2 Assessment of Cytochrome c Oxidase (COX; Complex IV) Activity in the Human Frozen Sections
3.3 Assessment of Succinate Dehydrogenase (SDH; Complex II) Activity in the Human Frozen Sections
3.4 Assessment of NADH Dehydrogenase (Diaphorase; Complex I) Activity in the Human Frozen Sections
4 Notes
References
Chapter 7: An Update on Isolation of Functional Mitochondria from Cells for Bioenergetics Studies
1 Introduction
2 Materials
2.1 Isolation of Crude Mitochondrial Fraction from Cell Lines
2.1.1 Solutions and Chemicals
2.1.2 Equipment and Accessories
2.2 Cell Culture
2.2.1 Materials and Accessories
2.2.2 Media and Solutions
2.3 Determination of Protein Concentration
2.3.1 Equipment, Accessories and Reagents
2.4 Measurement of the Mitochondrial Transmembrane Potential
2.4.1 Solutions and Reagents
2.4.2 Equipment and Accessories
2.5 Measurement of the Oxygen Consumption
2.5.1 Solutions and Reagents
2.5.2 Equipment
3 Methods
3.1 Isolation of Crude Mitochondrial Fraction from Cell Lines
3.2 Measurement of Protein Concentration
3.3 Measurement of the Mitochondrial Membrane Potential with the Use of Safranin O
3.4 Measurement of Oxygen Consumption (Respiration)
4 Notes
References
Chapter 8: PCR-Based Determination of Mitochondrial DNA Copy Number in Multiple Species
1 Introduction
2 Materials
2.1 DNA Isolation
2.1.1 C. elegans, Small Number
2.1.2 C. elegans, Large Numbers or Animal Tissue
2.1.3 Cultured Cells
2.2 DNA Quantification (See Note 1)
2.3 Real-Time PCR
2.4 Quantitative, Non-Real-Time, PCR
3 Methods
3.1 DNA Isolation
3.1.1 C. elegans, Small Number of Worms
3.1.2 C. elegans, Large Number of Worms or Animal Tissue (Skip Steps 1 and 2)
3.1.3 Cell Culture Samples
3.2 DNA Quantification
3.3 Real-Time PCR
3.3.1 C. elegans Samples with Standard Curve
3.3.2 All Other Real-Time PCR Samples, Without Standard Curve
3.3.3 Quantitative (NonReal-Time) PCR
3.4 Data Analysis
3.4.1 C. elegans and Human Samples, Real-Time PCR with Standard Curve
3.4.2 Other Species, Real-Time PCR Without Standard Curve
3.4.3 Quantitative, Non-real-Time PCR
4 Notes
References
Chapter 9: Methods to Monitor Mitophagy and Mitochondrial Quality: Implications in Cancer, Neurodegeneration, and Cardiovascul...
1 Introduction
1.1 Role of Mitophagy in Pathology
1.1.1 Mitophagy in CVD
1.1.2 Mitophagy in Cancer
1.1.3 Mitophagy in Neurodegenerative Diseases
2 Monitoring Mitophagy Using Fluorescent Probes
2.1 Colocalization Autophagosome and Mitochondria by Confocal Fluorescent Microscopy
2.1.1 Materials
2.1.2 Methods
2.1.3 Data Handling
2.1.4 Notes
2.2 Colocalization Parkin and Mitochondrial Marker
2.2.1 Materials
2.2.2 Methods
2.2.3 Data Handling
2.2.4 Notes
3 Monitoring Mitophagy Using Nonfluorescence Methods
3.1 Evaluation of Mitochondrial Protein Amounts
3.1.1 Materials
3.1.2 Method
3.1.3 Data Handling/Processing
3.1.4 Notes
3.2 Parkin Translocation by Subcellular Fractionation
3.2.1 Materials
3.2.2 Method
3.2.3 Data Handling/Processing
3.2.4 Notes
3.3 Detecting and Quantifying Mitophagy in Human Body Fluids
3.3.1 Materials
3.3.2 Methods
3.3.3 Data Analysis
3.3.4 Notes
4 Measuring the Mitochondrial Balance
4.1 Assessment of the Mitochondrial Structures
4.1.1 Materials
4.1.2 Methods
4.1.3 Data Handling/Processing
4.1.4 Notes
4.2 Assessing Changes in Mitochondrial Functioning by Immunoblot of Mitochondrial Complexes
4.2.1 Materials
4.2.2 Methods
4.2.3 Data Handling/Processing
4.2.4 Notes
4.3 Assessment of Mitochondrial Function Utilizing Seahorse Extracellular Flux
4.3.1 Materials
4.3.2 Method
4.3.3 Data Handling/Processing
4.3.4 Notes
4.4 Monitor Mitochondrial Turnover with MitoTimer
4.4.1 Materials
4.4.2 Methods
4.4.3 Data Handling
4.4.4 Notes
4.5 Monitor Mitochondrial Energetic Levels with Fluorescent Probes
4.5.1 Materials
4.5.2 Methods
4.5.3 Data Handling Analysis
4.5.4 Notes
References
Chapter 10: Analysis of Proapoptotic Protein Trafficking to and from Mitochondria
1 Introduction
2 Materials
2.1 Nuclear, Mitochondrial, and Cytosolic Fraction Isolation from Cultured Cells
2.2 Western Blotting
2.3 Immunoprecipitation
2.4 Immunocytochemistry
3 Methods
3.1 Nuclear, Mitochondrial, and Cytosolic Fraction Isolation from Cells in Culture
3.2 Western Blotting
3.2.1 SDS-Page
3.2.2 Wet/Tank Electrophoretic Transfer
3.2.3 Enzymatic Immunodetection
3.3 Immunoprecipitation
3.4 Immunocytochemistry
3.4.1 Cell Seeding for Morphological Studies
3.4.2 Staining with a Mitochondrial-Specific Fluorescent Dye (Option 1)
3.4.3 Staining with a Mitochondrial-Specific Fluorescent Protein (Option 2)
3.4.4 Fixation and Permeabilization
3.4.5 Immunolabeling
3.4.6 Mounting Coverslips
4 Notes
References
Chapter 11: Exploring Liver Mitochondrial Function by 13C-Stable Isotope Breath Tests: Implications in Clinical Biochemistry
1 Introduction
1.1 Studying Liver Function
1.2 General Features of Breath Tests (BTs)
1.3 Liver Mitochondria Function
1.4 Assessing Liver Mitochondrial Function by Breath Tests
2 Materials
2.1 General Information
2.2 13C-KICA BT
2.3 13C-Methionine BT
2.4 13C-Octanoate BT
3 Methods
4 Notes
References
Chapter 12: Protocols for Mitochondria as the Target of Pharmacological Therapy in the Context of Nonalcoholic Fatty Liver Dis...
Abbreviations
1 Introduction
1.1 Nonalcoholic Fatty Liver Disease (NAFLD)
1.2 Mechanisms of Free Fatty Acid (FFA) Accumulation in the Liver in Health and Lipotoxicity During Insulin Resistance Status
1.3 Focus on Mitochondrial Function in the Liver
2 Therapeutic Strategies in NAFLD
2.1 General Measures
3 Potential Strategies Effective also on Mitochondria in NAFLD
3.1 Rationale for Targeting Mitochondrial Function
3.2 Physical Exercise
3.3 Antidiabetic Drugs
3.4 Bile Acids
3.5 Antioxidant Agents
3.6 Mitotherapy
3.7 Miscellanea of Novel Agents
4 Summary
References
Chapter 13: The Use of Reactive Oxygen Species Production by Succinate-Driven Reverse Electron Flow as an Index of Complex 1 A...
1 Introduction
2 Materials
2.1 Preparation Steps for Isolation of Brown Adipose Tissue Mitochondria
2.2 Components for Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE)
2.3 Components for Immunoblotting
2.4 Molecular Weight Standards and Antibodies
2.5 Incubation Medium and Fluorimeter Used for Measuring Reactive Oxygen Species Production
2.6 Components for Complex 1 Assay
2.7 Components for Complex 2 Assay
3 Methods
3.1 Isolation of Brown Adipose Tissue Mitochondria
3.2 Measurement of Mitochondrial Reactive Oxygen Species (H2O2) Generation
3.3 SDS PAGE Electrophoresis
3.4 Immunoblotting
3.5 Complex I Activity Assay
3.6 Complex II Activity Assay
3.7 Statistical Analysis
4 Notes
References
Chapter 14: Mitochondrial Regulation Assessment by 13C-NMR Isotopomer Analysis
1 Introduction
2 OXPHOS Versus Aerobic Glycolysis
3 13C NMR Isotopomer Analysis
4 Protocols for NMR Evaluation of Mitochondrial Metabolic Regulation Using Cell Cultures
4.1 Untargeted Metabolomic Analysis of Cells by High-Rotation Magic-Angle Spinning (HRMAS)
4.2 Decipher the Glycolytic Versus OXPHOS Character of the Cell Line Using 13C-Enriched Substrates and 13C NMR Isotopomer Anal...
4.2.1 Lipidic and Aqueous Cellular Extracts for NMR Analyses
4.3 Contributions of Glutamine to Intermediary Cellular Metabolism
References
Chapter 15: Assays for Determination of Cellular and Mitochondrial NAD+ and NADH Content
1 Introduction
1.1 Cycling Assay Description
1.2 Isotopic Assay Description
2 Materials
2.1 Materials for Cellular NAD+ Extraction
2.2 Materials for Cellular NADH Isolation
2.3 Reaction Mixture Components
2.4 Fluorescent Reading
2.5 HPLC Isolation
2.6 Isotopic Label Dilution
3 Methods
3.1 Treatment and Harvest Protocol for Cellular NAD+ Content Measurement
3.2 Extraction of NAD+ from Cells and Animal Tissues
3.3 Cycling Assay
3.4 Data Analysis
3.5 NADH Measurement
3.6 Isotope-Labeled NAD+ Assay
4 Notes
References
Chapter 16: Following the Dynamism of the Mitochondrial Network in T Cells
1 Introduction
1.1 Mitochondrial Dynamics in Mammalian Cells
1.2 Mitochondria Shape in T Cell
1.3 Different Ways to Analyze Mitochondrial Dynamism
1.3.1 In Vitro Studies
1.3.2 Different Ways to Analyze Mitochondrial Dynamism: In Vivo Studies
2 Analysis of the Mitochondria Shape by Immunofluorescence
2.1 Material
2.2 Methods
2.3 Notes
3 Visualization of the Mitochondrial Network by Live Imaging
3.1 Material
3.2 Methods
3.3 Notes
References
Chapter 17: Measuring PGC-1α and Its Acetylation Status in Mouse Primary Myotubes
1 Introduction
2 Materials
2.1 Solutions and Media Formulations
2.2 Other Equipment and Supplies
3 Methods
3.1 Isolation of Mouse Primary Myoblasts
3.2 Infection with Adenovirus Encoding FLAG-PGC-1α and Empty Vector
3.3 Immunoprecipitation and Western Blot
4 Notes
References
Index
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Methods in Molecular Biology 2310
Carlos M. Palmeira Anabela P. Rolo Editors
Mitochondrial Regulation Methods and Protocols Second Edition
METHODS
IN
MOLECULAR BIOLOGY
Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK
For further volumes: http://www.springer.com/series/7651
For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.
Mitochondrial Regulation Methods and Protocols Second Edition
Edited by
Carlos M. Palmeira Department of Life Sciences, University of Coimbra, Coimbra, Portugal
Anabela P. Rolo Department of Life Sciences, University of Coimbra, Coimbra, Portugal
Editors Carlos M. Palmeira Department of Life Sciences University of Coimbra Coimbra, Portugal
Anabela P. Rolo Department of Life Sciences University of Coimbra Coimbra, Portugal
ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1432-7 ISBN 978-1-0716-1433-4 (eBook) https://doi.org/10.1007/978-1-0716-1433-4 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Illustration Caption: Image created by Madalena Palmeira. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.
Preface This book attempts to show different pathways that converge into the regulation of mitochondrial function. The book integrates mitochondria with other cellular components, discussing the dynamic properties of mitochondria with an emphasis on how these processes respond to signaling events and how they affect cellular metabolism. This second edition brings recent and more updated information, as well as new chapters with different perspectives. As before, this book is intended for the use of advanced undergraduates, graduates, postgraduates, and beginning researchers in the areas of molecular and cellular biology, biochemistry, and bioenergetics. 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 regulation. Coimbra, Portugal
Carlos M. Palmeira Anabela P. Rolo
v
Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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1 Isolation of Mitochondria from Liver and Extraction of Total RNA and Protein: Analyses of miRNA and Protein Expression . . . . . . . . . . . . . . . . . . . . 1 ˜ o, Pedro M. Borralho, Clifford J. Steer, Andre´ L. Sima Rui E. Castro, and Cecı´lia M. P. Rodrigues 2 Determination of Oxidative Phosphorylation Complexes Activities. . . . . . . . . . . . 17 ˜ o S. Teodoro, Ivo F. Machado, Carlos M. Palmeira, Joa and Anabela P. Rolo 3 BN-PAGE-Based Approach to Study Thyroid Hormones and Mitochondrial Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Elena Silvestri, Assunta Lombardi, Federica Cioffi, and Fernando Goglia 4 Monitoring Mitochondrial Function in Mouse Embryonic Stem Cells (mESCs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Bibiana Correia, Maria Ineˆs Sousa, Ana F. Branco, ˜ o Ramalho-Santos and Joa 5 Mitochondrial Functional Assessment in Mammalian Gametes Using Fluorescent Probes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57 Renata S. Tavares, Sara Escada-Rebelo, Maria M. Soares, Andreia F. Silva, Teresa Almeida-Santos, Sandra Amaral, ˜ o Ramalho-Santos Ana Paula Sousa, and Joa 6 Update on the Histoenzymatic Methods for Visualization of the Activity of Individual Mitochondrial Respiratory Chain Complexes in the Human Frozen Sections . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Mariusz R. Wieckowski, Maciej Pronicki, and Agnieszka Karkucinska-Wieckowska 7 An Update on Isolation of Functional Mitochondria from Cells for Bioenergetics Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Magdalena Lebiedzinska-Arciszewska, Lech Wojtczak, and Mariusz R. Wieckowski 8 PCR-Based Determination of Mitochondrial DNA Copy Number in Multiple Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Tess C. Leuthner, Jessica H. Hartman, Ian T. Ryde, and Joel N. Meyer 9 Methods to Monitor Mitophagy and Mitochondrial Quality: Implications in Cancer, Neurodegeneration, and Cardiovascular Diseases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Simone Patergnani, Massimo Bonora, Esmaa Bouhamida, Alberto Danese, Saverio Marchi, Giampaolo Morciano, Maurizio Previati, Gaia Pedriali, Alessandro Rimessi, Gabriele Anania, Carlotta Giorgi, and Paolo Pinton
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13
14
15
16 17
Contents
Analysis of Proapoptotic Protein Trafficking to and from Mitochondria . . . . . . . Ignacio Vega-Naredo, Gabriela Oliveira, Teresa Cunha-Oliveira, ˜ o, and Paulo J. Oliveira Teresa Serafim, Vilma A. Sarda Exploring Liver Mitochondrial Function by 13C-Stable Isotope Breath Tests: Implications in Clinical Biochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emilio Molina-Molina, Harshitha Shanmugam, Domenica Di Palo, Ignazio Grattagliano, and Piero Portincasa Protocols for Mitochondria as the Target of Pharmacological Therapy in the Context of Nonalcoholic Fatty Liver Disease (NAFLD). . . . . . . . . . . . . . . . Ignazio Grattagliano, Agostino Di Ciaula, Jacek Baj, Emilio Molina-Molina, Harshitha Shanmugam, Gabriella Garruti, David Q. -H. Wang, and Piero Portincasa The Use of Reactive Oxygen Species Production by Succinate-Driven Reverse Electron Flow as an Index of Complex 1 Activity in Isolated Brown Adipose Tissue Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrea Dlaskova´, Kieran J. Clarke, Mary F. Rooney, and Richard K. Porter Mitochondrial Regulation Assessment by 13C-NMR Isotopomer Analysis . . . . . Francisco X. Carvalho, Ba´rbara Guerra-Carvalho, Ivana Jarak, and Rui A. Carvalho Assays for Determination of Cellular and Mitochondrial NAD+ and NADH Content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yue Yang and Anthony A. Sauve Following the Dynamism of the Mitochondrial Network in T Cells . . . . . . . . . . . Arianna Di Daniele, Luca Simula, and Silvia Campello Measuring PGC-1α and Its Acetylation Status in Mouse Primary Myotubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ o A. Amorim and David A. Sinclair Joa
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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271 287
301 311
Contributors TERESA ALMEIDA-SANTOS • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; Reproductive Medicine Unit, Centro Hospitalar e Universita´rio de Coimbra, Coimbra, Portugal; Faculty of Medicine, Polo 3, University of Coimbra, Coimbra, Portugal SANDRA AMARAL • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; University of Coimbra, IIIUC- Institute for Interdisciplinary Research, Coimbra, Portugal JOA˜O A. AMORIM • Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for Biology of Aging Research, Harvard Medical School, Boston, MA, USA; Center for Neurosciences and Cell Biology of the University of Coimbra, Coimbra, Portugal; IIIUC – Institute of Interdisciplinary Research, University of Coimbra, Coimbra, Portugal GABRIELE ANANIA • Department of Medical Sciences, Section of General and Thoracic Surgery, University of Ferrara, Ferrara, Italy JACEK BAJ • Department of Anatomy, Medical University of Lublin, Lublin, Poland MASSIMO BONORA • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy PEDRO M. BORRALHO • Faculty of Pharmacy, Research Institute for Medicines (iMed. ULisboa), Universidade de Lisboa, Lisbon, Portugal; Medical Department, Novartis Oncology, Porto Salvo, Portugal ESMAA BOUHAMIDA • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy ANA F. BRANCO • CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal SILVIA CAMPELLO • Department of Biology, University of Rome Tor Vergata, Rome, Italy FRANCISCO X. CARVALHO • Department of Life Sciences, Faculty of Sciences and Technology, University of Coimbra, Coimbra, Portugal RUI A. CARVALHO • Department of Life Sciences, Faculty of Sciences and Technology, University of Coimbra, Coimbra, Portugal; REQUIMTE/LAQV, Group of Pharmaceutical Technology, Faculty of Pharmacy, University of Coimbra, Coimbra, Portugal RUI E. CASTRO • Faculty of Pharmacy, Research Institute for Medicines (iMed.ULisboa), Universidade de Lisboa, Lisbon, Portugal ` degli Studi del Sannio, FEDERICA CIOFFI • Dipartimento di Scienze e Tecnologie, Universita Benevento, Italy KIERAN J. CLARKE • School of Biochemistry & Immunology, Trinity Biomedical Sciences Institute, Trinity College Dublin, Dublin, Ireland BIBIANA CORREIA • Department of Life Sciences, University of Coimbra, Coimbra, Portugal; CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal TERESA CUNHA-OLIVEIRA • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal
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Contributors
ALBERTO DANESE • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy AGOSTINO DI CIAULA • Clinica Medica “A. Murri”, Department of Biomedical Sciences and Human Oncology, University of Bari Medical School, Bari, Italy ARIANNA DI DANIELE • Department of Biology, University of Rome Tor Vergata, Rome, Italy DOMENICA DI PALO • Clinica Medica “A. Murri”, Department of Biosciences and Human Oncology (DIMO), Policlinico Hospital, University of Bari Medical School, Bari, Italy ANDREA DLASKOVA´ • School of Biochemistry & Immunology, Trinity Biomedical Sciences Institute, Trinity College Dublin, Dublin, Ireland SARA ESCADA-REBELO • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; University of Coimbra, IIIUC- Institute for Interdisciplinary Research, Coimbra, Portugal GABRIELLA GARRUTI • Section of Endocrinology, Department of Emergency and Organ Transplantations, University of Bari “Aldo Moro” Medical School, Bari, Italy CARLOTTA GIORGI • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy ` degli Studi del FERNANDO GOGLIA • Dipartimento di Scienze e Tecnologie, Universita Sannio, Benevento, Italy IGNAZIO GRATTAGLIANO • Clinica Medica “A. Murri”, Department of Biomedical Sciences and Human Oncology (DIMO), Policlinico Hospital, University of Bari Medical School, Bari, Italy; Italian College of General Practitioners and Primary Care, Bari, Italy BA´RBARA GUERRA-CARVALHO • Department of Life Sciences, Faculty of Sciences and Technology, University of Coimbra, Coimbra, Portugal JESSICA H. HARTMAN • Nicholas School of the Environment, Duke University, Durham, NC, USA IVANA JARAK • REQUIMTE/LAQV, Group of Pharmaceutical Technology, Faculty of Pharmacy, University of Coimbra, Coimbra, Portugal AGNIESZKA KARKUCINSKA-WIECKOWSKA • Department of Pathomorphology, The Children’s Memorial Health Institute, Warsaw, Poland MAGDALENA LEBIEDZINSKA-ARCISZEWSKA • Nencki Institute of Experimental Biology, Warsaw, Poland TESS C. LEUTHNER • Nicholas School of the Environment, Duke University, Durham, NC, USA ` degli Studi di Napoli “Federico ASSUNTA LOMBARDI • Dipartimento di Biologia, Universita II”, Naples, Italy IVO F. MACHADO • Department of Life Sciences and Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal SAVERIO MARCHI • Department of Clinical and Molecular Sciences, Marche Polytechnic University, Ancona, Italy JOEL N. MEYER • Nicholas School of the Environment, Duke University, Durham, NC, USA EMILIO MOLINA-MOLINA • Clinica Medica “A. Murri”, Department of Biomedical Sciences and Human Oncology (DIMO), Policlinico Hospital, University of Bari Medical School, Bari, Italy GIAMPAOLO MORCIANO • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy
Contributors
xi
GABRIELA OLIVEIRA • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal PAULO J. OLIVEIRA • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal CARLOS M. PALMEIRA • Department of Life Sciences and Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal SIMONE PATERGNANI • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy GAIA PEDRIALI • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy; Maria Cecilia Hospital, GVM Care & Research, Cotignola, Italy PAOLO PINTON • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy RICHARD K. PORTER • School of Biochemistry & Immunology, Trinity Biomedical Sciences Institute, Trinity College Dublin, Dublin, Ireland PIERO PORTINCASA • Clinica Medica “A. Murri”, Department of Biomedical Sciences and Human Oncology (DIMO), Policlinico Hospital, University of Bari Medical School, Bari, Italy MAURIZIO PREVIATI • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy MACIEJ PRONICKI • Department of Pathomorphology, The Children’s Memorial Health Institute, Warsaw, Poland JOA˜O RAMALHO-SANTOS • Department of Life Sciences, University of Coimbra, Coimbra, Portugal; CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal ALESSANDRO RIMESSI • Department of Medical Sciences, Section of Pathology, Oncology and Experimental Biology, Laboratory for Technologies of Advanced Therapies (LTTA), University of Ferrara, Ferrara, Italy CECI´LIA M. P. RODRIGUES • Faculty of Pharmacy, Research Institute for Medicines (iMed. ULisboa), Universidade de Lisboa, Lisbon, Portugal ANABELA P. ROLO • Department of Life Sciences and Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal MARY F. ROONEY • School of Biochemistry & Immunology, Trinity Biomedical Sciences Institute, Trinity College Dublin, Dublin, Ireland IAN T. RYDE • Nicholas School of the Environment, Duke University, Durham, NC, USA VILMA A. SARDA˜O • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal ANTHONY A. SAUVE • Department of Pharmacology, Weill Cornell Medical College, New York, NY, USA TERESA SERAFIM • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal HARSHITHA SHANMUGAM • Clinica Medica “A. Murri”, Department of Biomedical Sciences and Human Oncology (DIMO), Policlinico Hospital, University of Bari Medical School, Bari, Italy
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Contributors
ANDREIA F. SILVA • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; University of Coimbra, IIIUC- Institute for Interdisciplinary Research, Coimbra, Portugal ` degli Studi del Sannio, ELENA SILVESTRI • Dipartimento di Scienze e Tecnologie, Universita Benevento, Italy ANDRE´ L. SIMA˜O • Faculty of Pharmacy, Research Institute for Medicines (iMed.ULisboa), Universidade de Lisboa, Lisbon, Portugal LUCA SIMULA • Department of Biology, University of Rome Tor Vergata, Rome, Italy DAVID A. SINCLAIR • Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for Biology of Aging Research, Harvard Medical School, Boston, MA, USA MARIA M. SOARES • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal ANA PAULA SOUSA • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; Reproductive Medicine Unit, Centro Hospitalar e Universita´rio de Coimbra, Coimbra, Portugal; Faculty of Medicine, Polo 3, University of Coimbra, Coimbra, Portugal MARIA INEˆS SOUSA • Department of Life Sciences, University of Coimbra, Coimbra, Portugal; CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal CLIFFORD J. STEER • Departments of Medicine, and Genetics, Cell Biology and Development, University of Minnesota Medical School, Minneapolis, MN, USA RENATA S. TAVARES • CNC-Center for Neuroscience and Cell Biology, CIBB, Polo 3, University of Coimbra, Coimbra, Portugal; University of Coimbra, IIIUC- Institute for Interdisciplinary Research, Coimbra, Portugal ˜ JOAO S. TEODORO • Department of Life Sciences and Center for Neurosciences and Cell Biology, University of Coimbra, Coimbra, Portugal IGNACIO VEGA-NAREDO • CNC - Center for Neuroscience and Cell Biology, UC-Biotech, University of Coimbra, Cantanhede, Portugal DAVID Q. -H. WANG • Division of Gastroenterology and Liver Diseases, Department of Medicine and Genetics, Marion Bessin Liver Research Center, Einstein-Mount Sinai Diabetes Research Center, Albert Einstein College of Medicine, Bronx, NY, USA MARIUSZ R. WIECKOWSKI • Laboratory of Mitochondrial Biology and Metabolism, Nencki Institute of Experimental Biology, Warsaw, Poland LECH WOJTCZAK • Nencki Institute of Experimental Biology, Warsaw, Poland YUE YANG • Department of Pharmacology, Weill Cornell Medical College, New York, NY, USA
Chapter 1 Isolation of Mitochondria from Liver and Extraction of Total RNA and Protein: Analyses of miRNA and Protein Expression Andre´ L. Sima˜o, Pedro M. Borralho, Clifford J. Steer, Rui E. Castro, and Cecı´lia M. P. Rodrigues Abstract Several studies have indicated the presence of microRNAs (miRNAs) within mitochondria although the origin, as well as the biological function, of these mitochondrially located miRNAs is largely unknown. The identification and significance of this subcellular localization is gaining increasing relevance to the pathogenesis of certain disease states. Here, we describe the isolation of highly purified mitochondria from rat liver by differential centrifugation, followed by RNAse A treatment to eliminate contaminating RNA. The coupled extraction of total RNA and protein is a more efficient design for allowing the downstream evaluation of miRNA and protein expression in mitochondria. Key words Differential centrifugation, miRNA, Mitochondria, Protein, TRIzol™
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Introduction Mitochondria are central organelles in the regulation of cellular homeostasis, playing a pivotal role in energy metabolism and cell viability, with their dysfunction or dysregulation being associated with multiple diseases. Human mitochondria harbor a compact circular genome of 16,569 bp in length, encoding 13 protein subunits of the electron transport chain, and a number of noncoding RNAs. Replication and transcription of mitochondrial DNA (mtDNA) are initiated from the small noncoding region known as the D loop and regulated by proteins encoded in the nuclear genome, which are posttranslationally imported into the mitochondria [1]. In addition, transcription and translation of mtDNA and mitochondrial transcript processing are regulated by several types of noncoding RNAs either encoded in the mitochondrial genome or in the nuclear genome following translocation to the mitochondria [2]. Interestingly, mitochondrial RNAs are transcribed from both
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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strands as long polycistronic precursor transcripts, which undergo processing and final release of both noncoding and coding RNAs, including tRNAs, rRNAs, and mRNAs. In the last decades, microRNAs (miRNAs) have emerged as a class of noncoding RNAs processed from endogenous transcripts, functioning as master regulators and fine-tuners of the genome, via posttranscriptional mechanisms. Mostly, miRNA gene-silencing occurs via miRNA-mRNA binding, typically with incomplete albeit high sequence complementarity between the 30 -untranslated region target sites of mRNA transcripts and the miRNA seed sequence located at position 2–8 of its 50 -end [3]. This allows miRNAs to regulate the expression of multiple target genes and signaling pathways involved in the regulation of key cellular processes including cell growth, proliferation, differentiation, and apoptosis [4], as well as mitochondrial function [2]. Consequently, deregulation of a given miRNA can lead to malfunction of pivotal cellular mechanisms, contributing to disease onset and/or progression. Therefore, it is not surprising that miRNA modulation is increasingly demonstrated as a relevant therapeutic strategy in human disease [5–8]. Since their discovery, miRNAs have been identified in multiple biological systems. Currently, more than 2600 mature human miRNAs have been identified (miRBase Version 22), which are predicted to regulate at least 60% of proteincoding genes. Other studies, including one by our group, have demonstrated the presence of mature miRNAs not only in the cytoplasm, where target gene-silencing of the mRNA transcript occurs, but also in other cellular compartments, including the nucleus [9], nucleolus [10], and mitochondria [11–14]. In addition to mature miRNAs, precursor miRNAs have also been found in mitochondria [13]. However, it is not known whether mitochondrial-localized miRNAs (also called mitomiRs) are encoded in the mitochondrial genome or in the nuclear genome and later imported into the mitochondria. Nevertheless, and regardless of their provenance from the nuclear or mitochondrial genome, the biological and pathophysiological implications of this unexpected finding are incompletely explored. It is conceivable that mitochondria-localized miRNAs may putatively function as posttranscriptional regulators, or fine-tuners, of the mitochondrial genome. This is illustrated by the mitochondrial translocation of both miR-1 and miR-181c, two cytoplasmatic mature miRNAs encoded in the nucleus. They function to regulate expression of mitochondria-encoded proteins such as cytochrome c oxidase subunit 1 (mt-COX1) and NADH: ubiquinone oxidoreductase core subunit 1 (mt-ND-1), in rodent myocytes [15, 16]. The first evidence that RNAi components may localize to the mitochondria was provided back in 2005, by the demonstration that human AGO2 interacts with mitochondrial tRNAmet [17]. Interestingly, miRNAs were later identified in mouse liver
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
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mitochondria by small RNA sequencing [18]. Aside from this unexpected finding, it was proposed that the presence of miRNAs in mouse liver mitochondria might arise from cytosolic contamination of the isolated mitochondria. Importantly, we have overcome this technical limitation by treating isolated and purified rat liver mitochondria with RNAse A, to ensure that mitochondria are devoid of contaminant RNAs from the cytosol or other cellular organelles and compartments. Coupling this relevant purification step to the detection of mature miRNAs by microarray, followed by northern blot confirmation, we have clearly demonstrated that mature miRNAs are indeed present in highly purified mitochondria [11]. This has been further confirmed in other studies of mitomiR regulation of crucial biological processes, particularly when the role of mitochondria is inevitable [19], although the cellular dynamics of intracellular miRNA translocation remains under debate. The identification of miRNAs in mitochondria and the exploration of their biological role in this subcellular organelle have been gaining increasing relevance. Mitochondria may be easily isolated and purified from multiple organs or cultured cells. The methodologies described in this chapter allow the straightforward isolation of highly purified mitochondria, and the coupled extraction of total RNA and protein, allowing downstream evaluation and comparison of mitochondrial miRNA and protein expression, in the same sample.
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Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water to attain a sensitivity of 18 MΩ cm at 25 C) and analytical grade reagents. Prepare and store all reagents at 4 C (unless otherwise indicated). Diligently follow all waste disposal regulations when disposing waste materials.
2.1 Mitochondrial Isolation: Materials and Reagents
1. Speed-Controlled Mechanical Drill: Tri-R model K41 skill drill (Tri-R Instruments, NY, USA). 2. Tissue Grinder: Glass mortar with radially serrated PTFE pestle (Fisher Scientific, NH, USA). 3. Ultraspin Buffer: 0.25 M sucrose, 1 mM ethylene glycol-bis (2-aminoethylether)- N,N,N0 ,N0 -tetraacetic acid (EGTA). Add 150 ml of water to a graduated cylinder. Weigh 21.39 g sucrose and 95.1 mg EGTA and transfer to the graduated cylinder. Mix and adjust pH to 7.4 with KOH. Complete volume with water to 250 mL (see Note 1). 4. Percoll (Colloidal PVP-coated silica for cell separation) (#P1644, Sigma-Aldrich, Missouri, USA).
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5. Homogenate Buffer: 70 mM sucrose, 220 mM mannitol, 1 mM EGTA, 10 mM 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid, N-(2-Hydroxyethyl) piperazine-N0 -(2-ethanesulfonic acid) (HEPES), pH 7.4. Add 700 mL of water to a glass beaker. Weigh 23.6 g sucrose, 40.8 g mannitol, 380.4 mg EGTA, and 2.38 g HEPES and transfer to the cylinder. Mix and adjust pH to 7.4 with KOH. Adjust volume to 1 L. 6. Mitochondria Wash Buffer 1: 0.1 M KCl, 5 mM 3-[N-Morpholino]propane sulfonic acid (MOPS), and 1 mM EGTA. Add 800 mL of water to a glass beaker. Weigh 7.46 g KCl, 1.05 g MOPS, and 0.380 mg EGTA and transfer to the glass beaker. Mix and adjust pH to 7.4 with KOH. Adjust volume to 1 L. 7. Mitochondria Wash buffer 2: 0.1 M KCl, 5 mM MOPS. Add 800 mL of water to a glass beaker. Weigh 7.46 g KCl and 1.05 g MOPS and transfer to the glass beaker. Mix and adjust pH to 7.4 with KOH. Adjust volume to 1 L. 8. Mitochondria Suspension Buffer: 125 mM sucrose, 50 mM KCl, 5 mM HEPES, and 2 mM KH2PO4. Add 70 mL of water to a glass beaker. Weigh 4.28 g sucrose, 372.8 mg KCl, 119.15 mg HEPES, and 27 mg KH2PO4 and transfer to the glass beaker. Mix and adjust volume to 100 mL. 9. EDTA-Free Complete®-Mini Protease Inhibitor Cocktail (#04693159001, Roche Applied Science, Penzberg, Germany). 10. Chelex™ 100 Resin (#143–2832, Bio-Rad Laboratories, CA, USA). 11. RNAse A, 20 mg/mL in 50 mM Tris–HCl (pH 8.0), 10 mM EDTA (#12091–039, Invitrogen™, Thermo Fisher Scientific, Massachusetts, USA). 2.2 Total RNA Isolation: Materials and Reagents
1. Nanodrop (Thermo Scientific). 2. TRIzol™ (#15596–026, Scientific).
Invitrogen™,
Thermo
Fisher
3. Chloroform. 4. Isopropanol. 5. UltraPure™ DNase/RNase-Free Distilled Water (#10977–035, Invitrogen™, Thermo Fisher Scientific). 6. 75% (vol/vol) Ethanol: Add 15 mL of ethanol and 5 mL of UltraPure™ DNase/RNase-Free Distilled Water in a sterile tube and mix well. Store at room temperature.
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
2.3 miRNA Expression Analysis: Materials and Reagents
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1. Thermal cycler. 2. miScript® II RT Kit (#218161, Qiagen, Hilden, Germany): miScript Reverse Transcriptase Mix, 10x miScript Nucleics Mix, and 5x miScript HiFlex Buffer. 3. UltraPure™ DNase/RNase-Free Distilled Water (#10977–035, Invitrogen™, Thermo Fisher Scientific). 4. miScript SYBR® Green PCR Kit (#218075, Qiagen). 5. QuantStudio™ 7 Flex Real-Time PCR System (or equivalent) (Applied Biosystems™, Thermo Fisher Scientific).
2.4 Total Protein Isolation: Materials and Reagents
1. Compact Ultrasonic Device: model UP100H—100 watts, ultrasonic frequency 30 kHz (Hielscher Ultrasonics GmbH, Teltow, Germany). 2. Tris–HCl 1 M, pH 8.0 (#T2694, Sigma-Aldrich). 3. Protein Wash Buffer: 0.3 M guanidine hydrochloride in 95% ethanol. Add 142.5 mL of ethanol and 7.5 mL of water to a graduated cylinder to prepare 150 mL of 95% ethanol. Add 80 mL of 95% ethanol to a beaker. Weigh 2.87 g of guanidine hydrochloride and transfer to the beaker. Bring the volume to 100 mL using 95% ethanol and mix well. Store at room temperature. 4. 1% (wt/vol) Sodium Dodecyl Sulfate (SDS): Add 80 mL of water to a graduated cylinder. Weigh 1 g of SDS and transfer to the cylinder. Mix well and bring the volume to 100 mL. Store at room temperature. 5. 8 M Urea in Tris–HCl 1 M, pH 8.0: Add 80 mL of water to a graduated cylinder. Weigh 48.05 g of urea and transfer to the cylinder. Mix well and bring the volume to 100 mL. Store at room temperature. 6. Protein Resuspension Buffer: 1:1 (vol/vol) of 1% SDS and 8 M urea in Tris–HCl 1 M, pH 8.0. Add 25 mL of 1% SDS solution and 25 ml of 8 M urea in Tris–HCl 1 M, pH 8.0 to a glass flask, as 50 mL of protein resuspension buffer. Mix well and store at room temperature.
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Methods
3.1 Mitochondria Isolation
The flowchart in Fig. 1 depicts the protocol followed after preparing the buffers and obtaining liver fragments. 1. Prepare two self-generating gradients (ultraspin buffer: Percoll (75:25); vol/vol) by adding 15 mL of ultraspin buffer and 5 mL of percoll to each ultracentrifuge tube. Mix well and centrifuge the tubes at 43,000 g for 30 min at 4 C. Carefully store the gradients at 4 C until use (see Note 2).
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Homogenize the samples as a 10% (wt/vol) homogenate, in homogenate buffer with protease inhibitor Use 6-10 up & down strokes with speed controlled mechanical drill and tissue grinder, at 800 rpm 600 x g, 10 min, 4ºC
Collect the supernatant (discard the pellet) 1,100 x g, 10 min, 4ºC
Collect the supernatant (discard the pellet) 7,700 x g, 10 min, 4ºC
Collect the pellet (discard the supernatant)
Resuspend the pellet in homogenate buffer with protease inhibitor Carefully and slowly layer on top of self-generating gradient 43,000 x g, 1 h, 4ºC
Collect the lower darker mitochondrial band Resuspend in mitochondria wash buffer 1 7,700 x g, 10 min, 4ºC
Wash purified mitochondria twice by resuspending in Chelex-100-treated mitochondria wash buffer 2 7,700 x g, 10 min, 4ºC
Collect the pellet (discard the supernatant)
Resuspend in Chelex-100-treated mitochondria suspension buffer Add 2 mg of RNAse A per mL Incubate 1 h, 37ºC 12,000 x g, 3 min, 4ºC
Collect the pellet (discard the supernatant)
Wash mitochondria pellet twice by resuspending in ChelexTM 100-treated mitochondria suspension buffer 12,000 x g, 3 min, 4ºC
Collect the mitochondrial pellet (discard supernatant) Proceed to RNA extraction (section 3.2.) (alternatively, store the pellet at -80ºC)
Fig. 1 Flowchart of mitochondrial isolation. Includes all steps following the preparation of buffers and collecting the liver sample for processing
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
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2. Prepare homogenate buffer containing EDTA-free Complete®-Mini protease inhibitor cocktail by adding 12 tablets to 120 mL of homogenate buffer. Mix to dissolution and place on ice (see Note 3). 3. Prepare 1% (wt/vol) Chelex™ 100-treated mitochondria wash buffer 2. Add 1.5 g of Chelex™ 100 to 150 mL of mitochondria wash buffer 2. Mix well by vigorous shaking by hand and incubate for 10 min at room temperature (see Note 4). Place on ice until use. 4. Prepare 1% (wt/vol) Chelex™ 100-treated mitochondria suspension buffer. Add 100 mg of Chelex™ to 10 mL of suspension buffer. Mix well by vigorous shaking by hand and incubate for 10 min at room temperature (see Note 4). Place on ice until use. 5. Fill three glass petri dishes with 50 mL of ice-cold homogenate buffer and place them on ice, together with an additional empty glass plate. 6. Sacrifice an adult male 175–200 g Sprague-Dawley or Wistar rat by exsanguination under CO2 anesthesia. 7. Remove the rat liver, and using tweezers, rinse the liver in the three petri dishes containing ice-cold homogenate buffer (prepared in step 5) and keep the rat liver on ice in the final rinse solution (see Note 5). 8. Using the ice-cold empty glass plate and a laboratory scale, weigh approximately 10 g of rat liver and mince the sample into small fragments using scissors (see Note 6). 9. Homogenize the sample as a 10% (wt/vol) homogenate. Homogenize the 10 g of minced liver in 100 mL ice-cold homogenate buffer containing EDTA-free Complete®-Mini protease inhibitor cocktail (prepared in step 2), using 6–10 complete up and down strokes with a speed-controlled mechanical drill and tissue grinder, at 800 rpm. Place the homogenate on ice. 10. Centrifuge the homogenate at 600 g for 10 min at 4 C. Collect the supernatant and discard the pellet (see Note 7). 11. Centrifuge the supernatant at 1100 g for 10 min at 4 C. Collect the supernatant and discard the pellet (see Note 8). 12. Centrifuge the supernatant at 7700 g for 10 min at 4 C. Discard the supernatant and keep the pellet (see Note 9). 13. Resuspend the pellet in 4 mL of homogenate buffer containing EDTA-free Complete®-Mini protease inhibitor cocktail (prepared in step 2), to obtain a crude mitochondrial extract. 14. Carefully, and slowly, layer 2 mL of the crude mitochondrial extract to each of the 20 mL of self-generating gradients (prepared in step 1). To purify the mitochondria, centrifuge at 43,000 x g for 1 h at 4 C.
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15. Carefully collect the lower yellowish-brown mitochondrial band and resuspend each in 30 mL of mitochondrial wash buffer 1. 16. Centrifuge at 7700 g for 10 min at 4 C. Discard the supernatant and keep the purified mitochondrial pellet. 17. Perform two sequential washes of the purified mitochondria by resuspending the mitochondrial pellet in 30 mL of Chelex™ 100-treated mitochondrial wash buffer 2 (prepared in step 3) (see Note 10), followed by centrifugation at 7700 g for 10 min at 4 C, discarding the supernatant and keeping the pellet. 18. Resuspend and pool the purified mitochondrial samples in 1–2 mL of Chelex™ 100-treated mitochondrial suspension buffer (prepared in step 4). 19. Perform RNAse treatment by adding 2 mg of RNAse A per mL of purified mitochondrial suspension and incubate at 37 C for 1 h (see Note 11). 20. Centrifuge the mitochondrial suspension at 12,000 g for 3 min at 4 C. Discard supernatant and maintain the purified mitochondrial pellet on ice. 21. Wash the mitochondrial pellet two times by resuspending in Chelex™ 100-treated mitochondrial suspension buffer (prepared in step 4) (see Note 10), and centrifuging at 12,000 g for 3 min at 4 C. 22. Discard the supernatant and place the purified mitochondria on ice. Proceed directly to total RNA extraction (go to Subheading 3.2) (see Note 12). 3.2 Total RNA Extraction from Isolated Mitochondria
The flowchart in Fig. 2 depicts the protocol followed. 1. Add TRIzol™ to the purified mitochondria pellet and pipette up and down to homogenize the sample (see Note 13). Incubate for 5 min at room temperature. 2. Add 0.2 mL of chloroform per mL of TRIzol™ reagent used for sample homogenization and vigorously shake by hand for 15 s, followed by 3 min incubation at room temperature. 3. Centrifuge at 12,000 g for 15 min at 4 C to allow phase separation and collect the upper aqueous phase into a new tube (see Note 14). The interphase and lower phenol-chloroform phase may be stored at 4 C, or at 80 C for prolonged storage, for subsequent total protein extraction (Subheading 3.4) (see Note 15). 4. Precipitate the RNA by adding 0.5 mL of isopropanol per mL of TRIzol™ reagent used for sample homogenization to the aqueous phase containing the RNA. Shake by hand for 15 s and incubate at room temperature for 10 min (see Note 16).
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
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Add TRIzolTM to the purified mitochondria pellet Homogenize the sample by pipetting up and down Incubate 5 min, room temperature
Add chloroform Vigorously shake by hand for 15 sec Incubate 3 min, room temperature 12,000 x g, 15 min, 4ºC
Collect the upper aqueous phase to a new tube Save interphase and lower phenol-chloroform phase at 4ºC or -80ºC for protein extraction (section 3.4)
Precipitate RNA by adding isopropanol to the aqueous phase Shake by hand for 15 sec Incubate 10 min, room temperature 12,000 x g, 10 min, 4ºC
Collect the pellet (discard the supernatant)
Wash RNA pellet with 75% ethanol Mix by inversion (10 times) 7,500 x g, 5 min, 4ºC
Collect the pellet (discard the supernatant) Air dry, 5 min
Resuspend RNA in DNAse/RNAse-free water Proceed to downstream applications or store at -80ºC
Fig. 2 Flowchart of total RNA extraction from isolated mitochondria
5. Centrifuge at 12,000 g for 10 min at 4 C and discard the supernatant. 6. Wash the RNA pellet by adding 1 mL of 75% ethanol per mL of TRIzol™ reagent used for sample homogenization and mix the tube by inversion ten times. 7. Centrifuge at 7500 g for 5 min at 4 C and discard the supernatant.
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8. Air-dry the RNA pellet for 5 min (see Note 17). 9. Resuspend the RNA in UltraPure™ DNase/RNase-Free Distilled Water and quantitate the RNA using NanoDrop 1000 (Thermo Scientific) (see Note 18). 10. Proceed to downstream applications or store the RNA at 80 C. 3.3 miRNA Expression Analysis
miRNA expression can be analyzed through quantitative reverse transcription PCR (qRT-PCR) using SYBR® Green (see Note 19). l
l
l
3.4 Total Protein Extraction from TRIzol™-Treated Isolated Mitochondria
Place sterile nuclease-free tubes on ice and add 1 μg of total RNA to 2 μL of 10x miScript Nucleics Mix, 2 μL miScript Reverse Transcriptase Mix, and 4 μL of 5x miScript HiFlex Buffer. Add DNase/RNase-Free distilled water up to 20 μL, vortex, and spin down the contents. Incubate for 60 min at 37 C, followed by 5 min at 95 C to inactivate reverse transcriptase and place on ice. Proceed to qRT-PCR or store the cDNA product at 20 C (see Note 20).
l
For qRT-PCR, prepare two independent reactions for each primer set, with a total volume of 25 μL containing 10–20 ng of cDNA, 2 QuantiTect SYBR Green PCR Master Mix, 10 miScript Universal Primer and 10 miScript Primer Assay of the specific miRNA primer.
l
Seal up the plate or tubes, spin down the content, and run the PCR using a QuantStudio™ 7 Flex Real-Time instrument (or equivalent). After 15 min at 95 C, SYBR green signal detection is performed for 40 cycles (each cycle: 15 s at 94 C, 30 s at 55 C, and 30 s at 70 C), with data collection at the end of every cycle.
l
Calculate the relative amounts of each miRNA based on the standard curve (see Note 21) and normalize to the level of 12S rRNA (see Note 22). Data may be presented as fold change against the respective control.
The flowchart in Fig. 3 depicts the protocol followed. 1. Start with the interphase and lower phenol-chloroform phase obtained after RNA isolation (step 3 of Subheading 3.2) and stored either at 4 C or at 80 C. If necessary, thaw the samples at room temperature and centrifuge at 12,000 g for 15 min at 4 C. Discard the remaining upper aqueous phase. 2. Add 0.3 mL of 100% ethanol per ml of TRIzol™ reagent used for sample homogenization to precipitate DNA. Mix by inversion and incubate for 3 min at room temperature.
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
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Fresh or frozen (-80ºC) interphase and lower phenolchloroform phase from total RNA extraction from isolated mitochondria (section 3.2) (Thaw frozen samples)
12,000 x g, 15 min, 4ºC
Discard remaining upper aqueous phase Precipitate DNA by adding 100% ethanol Mix by inversion Incubate 3 min, room temperature 2,000 x g, 5 min, 4ºC
Collect the phenol-ethanol supernatant (discard pellet) Precipitate protein with isopropanol Mix by inversion Incubate 10 min, room temperature 12,000 x g, 10 min, 4ºC
Collect the protein pellet (discard the supernatant)
Wash the protein pellet three times with protein wash buffer Vigorously shake by hand Incubate 20 min, room temperature 7,500 x g, 5 min, 4ºC
Wash protein pellet with 100% ethanol Vigorously shake by hand Incubate 20 min, room temperature 7,500 x g, 5 min, 4ºC
Solubilize the protein pellet in protein resuspension buffer Sonicate samples 3,200 x g, 10 min, 4ºC
Collect the clear supernatant containing the protein and store at -80º C for downstream applications
Fig. 3 Flowchart of total protein extraction from TRIzol™-treated isolated mitochondria
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3. Centrifuge at 2000 g for 5 min at 4 C. Transfer the phenolethanol supernatant to a new tube for protein isolation (see Note 23). 4. Add 1.5 mL of isopropanol per mL of TRIzol™ reagent used for sample homogenization. Mix by inversion and incubate for 10 min at room temperature. 5. Centrifuge at 12,000 g for 10 min at 4 C to precipitate protein and discard the supernatant. 6. Perform three sequential washes of the protein pellets by adding 2 mL of protein wash buffer per mL of TRIzol™ reagent used for sample homogenization, vigorously shaking the tubes by hand, incubating at room temperature for 20 min and centrifuging at 7500 g for 5 min at 4 C (see Note 24). 7. After the third wash and spin, add 2 mL of 100% ethanol per mL of TRIzol™ reagent, incubate at room temperature for 20 min, and centrifuge at 7500 g for 5 min at 4 C (see Note 24). Discard the supernatant and maintain the protein pellet on ice (see Note 25). 8. Solubilize the protein pellet by adding protein resuspension buffer, followed by five cycles of 15 s sonication and 30 s ice incubation, using a compact ultrasonic device with amplitude adjusted to 80% and pulse to 90%. 9. Centrifuge at 3200 g for 10 min at 4 C to sediment insoluble material. Discard the pellet and collect the supernatant containing the solubilized proteins to a fresh tube and store at 80 C for downstream applications.
4
Notes 1. Sucrose solutions are easily contaminated, even when stored at 4 C. Use the solution within 1 week. Ideally, prepare before use. 2. The gradients should always be prepared prior to use and carefully handled to avoid mixing and discontinuities that may alter the efficiency of the separations. 3. EDTA-free Complete®-Mini protease inhibitor cocktail should be added before use and the buffer kept on ice. Always prepare fresh, discarding remainder of the solution, to ensure maximal protease inhibition. 4. Treatment of buffers with chelating resin Chelex™ 100 should be performed before use. Discard the remainder of Chelex™ 100-treated buffers to ensure optimal clearing of the buffers. 5. Rinsing in three consecutive changes of homogenate buffer eliminates blood and other contaminants, which may be located on the outer surface of the liver during animal sacrifice
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . .
13
and organ collection. After the final rinse of the liver in ice-cold homogenate buffer, the buffer should be clear and devoid of blood and macroscopic debris. If necessary, perform additional rinses in similar fashion. 6. Additional liver fragments may be stored for subsequent analysis (e.g., protein and/or RNA extraction from whole liver) by snap-freezing the samples in liquid nitrogen, followed by storage at 80 C. 7. This step clears the sample from insoluble tissue, incompletely lysed cells and other debris. 8. This step further separates the sample from insoluble tissue, incompletely lysed cells and other debris. 9. This step precipitates mitochondria and other cellular organelles. 10. The Chelex™ 100 resin precipitates by gravity. Following the 10 min incubation at room temperature, the Chelex™ 100 resin deposits at the bottom of the solution. Pipette the buffer without disturbing the Chelex™ 100 resin deposit. 11. RNAse treatment of isolated mitochondria eliminates RNAs in solution and those located on the outer surface of the mitochondria, ensuring the absence of nonmitochondrial contaminating RNAs in the sample. Optionally, before and/or after RNAse A treatment, you may take a sample of purified mitochondria to evaluate mitochondria morphology and purity by transmission electron microscopy and western blot, as previously described [11]. 12. You may optionally snap-freeze the purified mitochondria in liquid nitrogen, followed by storage at 80 C. 13. Isolation of total RNA using TRIzol™ allows the isolation of microRNAs, suitable for multiple downstream applications [5– 7, 20–22]. Alternatively, samples may be homogenized in TRIzol™ reagent using a motor-driven Biovortexer (No1083; Biospec Products, Bartlesfield, OK) and disposable RNAse/DNAse-free sterile pestles (Thermo Fisher Scientific, Inc., Chicago, IL) [21]. 14. The sample mixture separates into an upper colorless aqueous phase, an interphase, and a lower red phenol-chloroform phase. The RNA containing the miRNAs is exclusively in the upper colorless aqueous phase. 15. Phenol-chloroform phases may be stored at 4 C overnight or at 80 C for long-term storage, of at least 2 years [23]. 16. Alternatively, you may incubate the samples at 20 C overnight to increase RNA precipitation or as a stop point in the protocol. 17. Do not allow the RNA pellet to completely dry as this reduces its solubility.
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18. Alternatively, you can quantitate the RNA by diluting the sample in UltraPure™ DNase/RNase-Free Distilled Water and determining its absorbance at 260 nM and 280 nM. RNA concentration may be determined using the formula: A260 dilution 40 ¼ μg of RNA per mL. 19. miRNA expression levels may also be determined by microarray analysis [11, 12, 20], northern blotting [22], Taqman RealTime RT-PCR [5–7, 21], or deep sequencing [24]. 20. cDNA can be kept at 4 C or at 20 C for up to 1 week. Alternatively, it may be kept at 80 C for long-term storage. 21. You may perform a standard curve for each miRNA/endogenous reference primer, doing a set of serial dilutions (1/1; 1/3; 1/9; 1/27; 1/81) of a cDNA pool from all samples used in the study. 22. For analysis of 12S rRNA expression, use the QuantiTect Primer Assay instead of the 10 miScript Primer Assay and do not use the 10 miScript Universal Primer. Other mitochondrial-specific genes, such as 16S rRNA, may also be used for PCR normalization. 23. This step precipitates DNA, with the proteins remaining in the phenol-ethanol supernatant. 24. It is not necessary to resuspend the protein pellet. Vigorously shake by hand and incubate at room temperature. Following centrifugation, completely remove each buffer solution to maximize wash efficacy. 25. Alternatively, the protein pellets may be snap-frozen in liquid nitrogen and stored at 80 C until resuspension.
Acknowledgments CMPR is supported by the EU H2020 Marie Sklodowska-Curie Project Foie Gras (grant 722619) and by FCT and FEDER (grant PTDC/MEDFAR/29097/2017 e LISBOA-01-0145-FEDER029097). References 1. Mercer TR, Neph S, Dinger ME, Crawford J, Smith MA, Shearwood AM, Haugen E, Bracken CP, Rackham O, Stamatoyannopoulos JA, Filipovska A, Mattick JS (2011) The human mitochondrial transcriptome. Cell 146 (4):645–658 2. Geiger J, Dalgaard LT (2017) Interplay of mitochondrial metabolism and microRNAs. Cell Mol Life Sci 74(4):631–646
3. Gebert LFR, MacRae IJ (2019) Regulation of microRNA function in animals. Nat Rev Mol Cell Biol 20(1):21–37 4. Lui PY, Jin DY, Stevenson NJ (2015) MicroRNA: master controllers of intracellular signaling pathways. Cell Mol Life Sci 72 (18):3531–3542 5. Castro RE, Ferreira DM, Afonso MB, Borralho PM, Machado MV, Cortez-Pinto H,
Isolation of Mitochondria from Liver and Extraction of Total RNA and. . . Rodrigues CM (2013) miR-34a/SIRT1/p53 is suppressed by ursodeoxycholic acid in the rat liver and activated by disease severity in human non-alcoholic fatty liver disease. J Hepatol 58 (1):119–125 6. Gomes SE, Simoes AE, Pereira DM, Castro RE, Rodrigues CM, Borralho PM (2016) miR-143 or miR-145 overexpression increases cetuximab-mediated antibody-dependent cellular cytotoxicity in human colon cancer cells. Oncotarget 7(8):9368–9387 7. Gomes SE, Pereira DM, Roma-Rodrigues C, Fernandes AR, Borralho PM, Rodrigues CMP (2018) Convergence of miR-143 overexpression, oxidative stress and cell death in HCT116 human colon cancer cells. PLoS One 13(1): e0191607 8. Zhang T, Hu J, Wang X, Zhao X, Li Z, Niu J, Steer CJ, Zheng G, Song G (2019) MicroRNA-378 promotes hepatic inflammation and fibrosis via modulation of the NF-κB-TNFα pathway. J Hepatol 70(1):87–96 9. Sarshad AA, Juan AH, Muler AIC, Anastasakis DG, Wang X, Genzor P, Feng X, Tsai PF, Sun HW, Haase AD, Sartorelli V, Hafner M (2018) Argonaute-miRNA complexes silence target mRNAs in the nucleus of mammalian stem cells. Mol Cell 71(6):1040–1050.e8 10. Reyes-Gutierrez P, Ritland Politz JC, Pederson T (2014) A mRNA and cognate microRNAs localize in the nucleolus. Nucleus 5 (6):636–642 11. Kren BT, Wong PY, Sarver A, Zhang X, Zeng Y, Steer CJ (2009) MicroRNAs identified in highly purified liver-derived mitochondria may play a role in apoptosis. RNA Biol 6 (1):65–72 12. Bian Z, Li LM, Tang R, Hou DX, Chen X, Zhang CY, Zen K (2010) Identification of mouse liver mitochondria-associated miRNAs and their potential biological functions. Cell Res 20(9):1076–1078 13. Barrey E, Saint-Auret G, Bonnamy B, Damas D, Boyer O, Gidrol X (2011) Pre-microRNA and mature microRNA in human mitochondria. PLoS One 6(5):e20220 14. Jagannathan R, Thapa D, Nichols CE, Shepherd DL, Stricker JC, Croston TL, Baseler WA, Lewis SE, Martinez I, Hollander JM (2015) Translational regulation of the mitochondrial genome following redistribution of mitochondrial microRNA in the diabetic heart. Circ Cardiovasc Genet 8(6):785–802 15. Zhang X, Zuo X, Yang B, Li Z, Xue Y, Zhou Y, Huang J, Zhao X, Zhou J, Yan Y, Zhang H, Guo P, Sun H, Guo L, Zhang Y, Fu XD (2014) MicroRNA directly enhances mitochondrial translation during muscle differentiation. Cell 158(3):607–619
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16. Das S, Kohr M, Dunkerly-Eyring B, Lee DI, Bedja D, Kent OA, Leung AK, Henao-Mejia J, Flavell RA, Steenbergen C (2017) Divergent effects of miR-181 family members on myocardial function through protective cytosolic and detrimental mitochondrial microRNA targets. J Am Heart Assoc 6(3):e004694 17. Maniataki E, Mourelatos Z (2005) Human mitochondrial tRNAMet is exported to the cytoplasm and associates with the Argonaute 2 protein. RNA 11(6):849–852 18. Lung B, Zemann A, Madej MJ, Schuelke M, Techritz S, Ruf S, Bock R, Huttenhofer A (2006) Identification of small non-coding RNAs from mitochondria and chloroplasts. Nucleic Acids Res 34(14):3842–3852 19. Song R, Hu X-Q, Zhang L (2019) Mitochondrial miRNA in cardiovascular function and disease. Cell 8(12):1475 20. Sarver AL, French AJ, Borralho PM, Thayanithy V, Oberg AL, Silverstein KA, Morlan BW, Riska SM, Boardman LA, Cunningham JM, Subramanian S, Wang L, Smyrk TC, Rodrigues CM, Thibodeau SN, Steer CJ (2009) Human colon cancer profiles show differential microRNA expression depending on mismatch repair status and are characteristic of undifferentiated proliferative states. BMC Cancer 9:401 21. Borralho PM, Simoes AE, Gomes SE, Lima RT, Carvalho T, Ferreira DM, Vasconcelos MH, Castro RE, Rodrigues CM (2011) miR-143 overexpression impairs growth of human colon carcinoma xenografts in mice with induction of apoptosis and inhibition of proliferation. PLoS One 6(8):e23787 22. Castro RE, Ferreira DM, Zhang X, Borralho PM, Sarver AL, Zeng Y, Steer CJ, Kren BT, Rodrigues CM (2010) Identification of microRNAs during rat liver regeneration after partial hepatectomy and modulation by ursodeoxycholic acid. Am J Physiol Gastrointest Liver Physiol 299(4):G887–G897 23. Simoes AE, Pereira DM, Amaral JD, Nunes AF, Gomes SE, Rodrigues PM, Lo AC, D’Hooge R, Steer CJ, Thibodeau SN, Borralho PM, Rodrigues CM (2013) Efficient recovery of proteins from multiple source samples after TRIzol® or TRIzol®LS RNA extraction and long-term storage. BMC Genomics 14:181 24. Sripada L, Tomar D, Prajapati P, Singh R, Singh AK (2012) Systematic analysis of small RNAs associated with human mitochondria by deep sequencing: detailed analysis of mitochondrial associated miRNA. PLoS One 7(9): e44873
Chapter 2 Determination of Oxidative Phosphorylation Complexes Activities Joa˜o S. Teodoro, Ivo F. Machado, Carlos M. Palmeira, and Anabela P. Rolo Abstract Mitochondria possess a genome that codes for proteins, in the same fashion as the nuclear genome. However, the small, circular mitochondrial DNA (mtDNA) molecule has a reduced base pair content, for it can only code for 2 rRNA, 22 tRNA molecules, and 13 proteins, all of them part of the mitochondrial respiratory chain. As such, all of the other mitochondrial components derive from nuclear genome. This separation leads to a requirement for a well-tuned coordination between both genomes, in order to produce fully functional mitochondria. A vast number of pathologies have been demonstrated to involve, to some extent, alterations in mitochondrial function that, no doubt, can be caused by alterations to the respiratory chain activity. As such, several methods and techniques have been developed to assess both content and function of mitochondrial proteins, in order to help understand mitochondrial involvement on the pathogenesis of disease. In this chapter, we will address some of these methods, with the main focus being on isolated mitochondria. Key words Mitochondria, Respiratory chain, Polarography, Mitochondrial protein complexes, Spectrophotometry
1
Introduction Mitochondria are the eukaryotic cell’s main site of ATP generation, accounting for roughly 95% of all ATP consumed in the cell. Along with ATP generation, mitochondria are also responsible for the storage of various molecules and ions (as, for example, Ca2+). It is also here, in a physically separated (from the cytosol) environment that several biochemical reactions can take place, namely the lipid β-oxidation, the citric acid cycle (or Krebs cycle), to name a few. As such, it comes as no surprise that mitochondrial data are extremely important to a variety of studies. The oxidative phosphorylation system is made up from several protein complexes (I–IV), which together with the mobile proteins ubiquinone (also known as coenzyme Q10) and cytochrome c make the respiratory chain, across the mitochondrial inner membrane.
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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This system takes the electrons from metabolic substrates and transports them in increasingly free-energy favorable electronic jumps toward O2, forming H2O at Complex IV. Since this electronic transport is energetically favorable, a lot of energy is released. These protein complexes (except Complex II) are assembled in such a way that this energy is harnessed to eject protons from the mitochondrial matrix, across the proton-impermeable inner membrane into the intermembrane space. This proton gradient can then be used by the 5th complex, the ATPSynthase to drive the phosphorylation of ADP into ATP. The respiratory chain protein complexes constituents are coded in both mitochondrial and nuclear genomes, and mutations on these genes are among the most common reasons for mitochondrial diseases. Other causes for diseases that range from diabetes to neurological diseases and aging involve the posttranscriptional modulation of mitochondrial respiratory activity [1]. As such, careful, reliable, and sensitive evaluation of the mitochondrial respiratory capacity is truly necessary to trace the pathological pathways and targets that characterize a disease. Here, we describe one protocol for the characterization of the activity of all these protein complexes in isolated mitochondria from tissue homogenates from rat livers. For other tissues, minute alterations can be performed to maximize yield and function.
2
Materials
2.1 Reagents and Buffers
Standard hepatic mitochondrial homogenization buffer: l
Sucrose 250 mM.
l
EGTA 0.5 mM.
l
Bovine serum albumin 0.5%.
l
HEPES 10 mM, pH 7.4.
Standard hepatic mitochondrial wash buffer: l
Sucrose 250 mM.
l
HEPES 10 mM, pH 7.4.
Standard hepatic mitochondrial respiratory buffer: l
Sucrose 130 mM.
l
KCl 50 mM.
l
MgCl2 5 mM.
l
KH2PO4 5 mM.
l
HEPES 10 mM, pH 7.4.
Determination of Oxidative Phosphorylation Complexes Activities
19
Standard ATPase reaction buffer: l
Sucrose 125 mM.
l
KCl 65 mM.
l
MgCl2 2.5 mM.
l
HEPES 50 mM, pH 7.2.
Biuret Reagent: To make Biuret reagent, weight 1.5 g of cupric sulfate pentahydrate (CuSO4 5.H2O) and dissolve it in 500 mL of water. Then, weight 6 g of sodium potassium tartrate tetrahydrate (NaKC4H4O6 4.H2O) and slowly add it to the dissolved cupric sulfate solution (see Note 1). Add 300 mL of a 10% solution of sodium hydroxide (NaOH). Add water to a final volume to 1 L, store in a light-protected glass bottle. It should be stable for a year. Molybdate Reagent: To produce Molybdate reagent, dissolve 5 g of ferrous sulfate (FeSO4) in 60 mL of water. After which, add 10 mL of a 10% ammonium molybdate ((NH4)6Mo7O24.4H2O) solution. Adjust the volume to 100 mL with water. Reagents: l
2,6-dichloroindophenol (DCIP).
l
5,500 -dithiobis-2-nitrobenzoic acid (DTNB).
l
Acetyl-CoA.
l
Antimycin A.
l
Ascorbate.
l
ATP.
l
Bovine serum albumin (BSA).
l
Decylubiquinone.
l
Deoxycholic acid (DOC).
l
HCl.
l
KCN.
l
KH2PO4.
l
KHCO3.
l
Lithium borohydride.
l
NADH.
l
Oxaloacetate.
l
Oxidized cytochrome c.
l
Phenazine methosulfate.
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2.2 Equipment Required
3
l
Rotenone.
l
Succinate.
l
Tetramethyl-phenylenediamine (TMPD).
l
Trichloroacetic acid (TCA).
l
Tris.
l
Triton X-100.
l
Ultrapure water.
l
50 mL centrifuge tubes.
l
Animal (rat, mice, and others).
l
Clark-type oxygen electrode, reaction chamber, and associated register.
l
Eppendorf-like 1.5 mL tubes.
l
Glass Potter-Elvehjem homogenizer with Teflon pestle.
l
Laboratory chronometer.
l
Magnetic stirrer and magnetic stir bar.
l
Orbital vortex.
l
Plastic cuvettes.
l
Power drill.
l
Precision pipettes.
l
Refrigerated centrifuge.
l
Small volume beakers.
l
Smooth paintbrushes.
l
Spectrophotometer with cuvette access.
l
Surgical scissors.
l
Surgical tweezers.
l
Test-tube holder.
l
Test tubes.
l
Water bath, preferably with lid.
l
Take the animal, starved overnight, and quickly sacrifice it by cervical dislocation and decapitation. Bleed the animal into the sink for 5–10 s.
l
Using surgical scissors and tweezers, cut open the animal’s abdomen, right below the ribcage, and remove the liver, in large pieces, into a beaker containing ice-cold homogenization
Methods
3.1 Isolation of Hepatic Mitochondria
Determination of Oxidative Phosphorylation Complexes Activities
21
buffer (see Note 2). Remove any adhering fat, fibrous tissue, or blood vessels from the liver chunks and thinly chop the liver into small pieces (see Note 3). Replace buffer.
3.2 Mitochondrial Quantification
l
Add approximately 5–6 mL of ice-cold homogenization buffer to each gram of chopped liver (typically 50 mL) and transfer this mix into a precooled glass Potter-Elvehjem homogenizer.
l
Homogenize the tissue using 3–4 up/down strokes of pestle rotating at roughly 300 rpm (see Note 4).
l
When the tissue is homogenized, transfer the homogenate into two previously cooled centrifuge tubes, balancing them with homogenization buffer. Centrifuge the homogenate at 800 g for 10 min at 4 C (see Note 5).
l
Carefully decant the supernatant into new cooled centrifugation tubes (see Note 6). Balance them with homogenization buffer and centrifuge at 10000 g for 10 min at 4 C.
l
Discard the supernatant as completely as possible and gently resuspend the pellet in a small volume (roughly 3–5 mL) of wash buffer (see Note 7).
l
Repeat the last step two more times. After the final centrifugation (3rd at 10,000 g, 4th total), resuspend the mitochondrial pellet into a small volume of wash buffer (see Note 8).
l
Let the mitochondria rest for 15–30 min (see Note 9).
l
During the mitochondrial rest period described above, it is possible to use this time to quantify the mitochondrial protein content, since all assays are normalized against the mitochondrial protein content. For isolated rat mitochondria, we recommend the use of the Biuret assay [2]. Briefly, follow Table 1. Tubes 0–3 are for the standard curve.
l
Prepare the tubes as in Table 1 (see Note 10). Mix the tubes in an orbital vortex and put them in a water bath, light-protected at 37 C for 5 min.
l
When the 5 min are over, measure the absorbance of the tubes’ content in standard plastic 4 mL cuvettes at 540 nm in a standard spectrophotometer (see Note 11). Construct a standard curve from the 1–3 tubes.
l
Calculate the average absorbance of your duplicates (see Note 12) and insert it the following formula: X ¼ ððAbs InsÞ=SlopeÞ DF, where Abs is the average absorbance of your samples, Ins is the standard curve intersection, Slope is the standard curve slope, and DF is the dilution factor. X is your sample’s protein content, expressed in mg/mL (see Note 13).
Joa˜o S. Teodoro et al.
22
Table 1 Biuret Assay preparation H2O Tube (μL)
BSA 0.4% (μL)
Wash buffer (μL)
Sample (μL)
DOC 10% (μL)
Biuret reagent (mL)
Protein content (mg)
0
500
0
50
–
50
2
0.0
1
250
250
50
–
50
2
1.0
2
125
375
50
–
50
2
1.5
3
0
500
50
–
50
2
2.0
X1
500
–
–
50
50
2
X
X2
500
–
–
50
50
2
X
Abbreviations are as follows: BSA bovine serum albumin, DOC deoxycholic acid
3.3 Citrate Synthase Activity Assay
Citrate synthase is a mitochondrial matrix enzyme. This assay is used as an indicator of mitochondrial numbers in tissue homogenates and impure mitochondrial preparations, as it has not been found to be deficient in any disease states. Respiratory chain enzyme activities can be expressed as citrate synthase ratios to correct for any variations in mitochondrial numbers in these preparations. Also, mitochondrial integrity can be assessed using fresh mitochondria by this assay by citrate synthase latency in the presence of Triton X-100. This assay monitors the release of free coenzyme A (CoA) from acetyl-CoA after the citrate synthase reactions initiated by the addition of oxaloacetate. This is achieved by Ellman’s reagent (5,500 -dithiobis-2-nitrobenzoic acid, DTNB) reaction with the free thiol groups of CoA and registering the absorbance at 412 nm. 1. Prepare the following reagents: (a) Tris 200 mM, pH 8.0. (b) Acetyl-CoA 100 mM. (c) DNTB 100 mM (4 mg/mL)—add a pinch of KHCO3 to help dissolve. (d) Oxaloacetate 10 mM (1.4 mg/mL)—add Tris to pH 7.0. (e) Triton X-100 10%. 2. For each sample, the reaction mix is made in a cuvette from 500 μL Tris, 20 μL acetyl-CoA, 20 μL DNTB, and 50 μg of mitochondrial protein. Add water to make 1 mL. After a light mix, record a stable baseline at 412 nm. 3. Start the reaction by adding 10 μL of oxaloacetate (see Note14). 4. Finally, add 0.1% (v/v—final concentration) Triton X-100 after recording the oxaloacetate rate (see Note 15).
Determination of Oxidative Phosphorylation Complexes Activities
23
With fresh mitochondria, the addition of oxaloacetate will cause an increase in the absorbance rate as intense as more citrate synthase is in the medium, meaning, the higher the rate, the more mitochondria are damaged. Adding Triton releases all of the citrate synthase still trapped inside mitochondria, which should demonstrate a steady value between preparations. The extinction coefficient for DTNB at 412 nm is of 13.6 M/ cm, which means that citrate synthase activity is calculated as follows: Activity ðnmol min mLÞ ¼ ðΔAbs Vt 1, 000, 000Þ= EC Δtime mg C½S , where ΔAbs is the change in absorbance, Vt is the final cuvette volume in microliters, mg is the mitochondrial protein content (for this example, it is 0.05), EC is the extinction coefficient (in this example, 13.6), C[S] is the sample concentration, and Δtime is the elapsed time between stable recordings of absorbance. 3.4 Individual Respiratory Chain Enzymatic Activities
Figure 1 indicates where and how these methods evaluate the different respiratory complexes. Below there are individual subheadings with the respective methods and techniques.
3.4.1 Complex I (NADH: Ubiquinone Oxidoreductase)
Complex I is the first protein complex of the mitochondrial oxidative phosphorylation system. It receives electrons from NADH (which is oxidized to NAD+) and transports them to the oxidized coenzyme-Q10 or ubiquinone. To this oxidation of NADH and electron transport to ubiquinone is coupled the vectorial ejection of protons from the mitochondrial matrix to the intermembrane space, creating a protonic gradient with an electric and pH component, which is used by the ATPSynthase to drive the phosphorylation of ADP into ATP. It is the largest and heaviest complex of the respiratory chain, and the deficiency in Complex I is probably the most frequently encountered cause of mitochondrial disease, and various mutations in the genes coding for both the nuclear and mitochondrially encoded subunits have already been described [3]. Here, we describe a method first published by Janssen and collaborators [4]. As the authors describe, this method relies on Complex I oxidizing NADH and giving the electrons to the artificial substrate decylubiquinone, which in turn delivers them to DCIP. It is then possible to spectrophotometrically follow the reduction of DCIP by measuring the absorbance at 600 nm. It is a highly sensitive method, since decylubiquinone does not accept electrons from other sources [5]. This method also has the advantage of not requiring an UV-light source, for by following the disappearance rate of NADH as described by other methods [1], one would require UV-ready cuvettes and a spectrophotometer with an UV lamp, which is harder and much more expensive to obtain. As such, recording at 600 nm, within the visible spectra, one can bypass such issues.
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Joa˜o S. Teodoro et al.
Fig. 1 Determination of the enzymatic activities of individual respiratory chain complexes. (a) Complex I activity is determined by exploiting its capacity of NADH oxidation in a process that releases electrons to be transferred to decylubiquinone and consequently to DCIP, leading to its reduction, which is possible to be spectrophotometrically assessed (see Subheading 3.4.1); (b) to evaluate complex II activity, PMS is utilized as an acceptor of electrons originating from succinate. PMS donates electrons to O2, and the oxygen consumption rate is assessed (see Subheading 3.4.2); (c) Complex III activity is spectrophotometrically evaluated by following the reduction of cytochrome c (see Subheading 3.4.3); (d) Complex IV activity is measured by following oxygen consumption rate after TMPD is used as an electron donor (see Subheading 3.4.4); and (e) ATP synthase activity is measured by taking advantage of its reverse ATPase function. The reaction between molybdate and Pi, originated from ATP hydrolysis, is spectrophotometrically assessed (see Subheading 3.4.5) l
Prepare respiration buffer (as described in Subheading 2.1).
l
To this solution, BSA 0.4% should be added to maximize Complex I activity [4]. Also add KCN 240 μM, antimycin A 4 μM, DCIP 60 μM, and decylubiquinone 70 μM. Prepare two tubes per sample, one with and other without rotenone 2 μM (see Note 16).
l
Add 0.3 mg of freeze-thawed mitochondrial protein to both tubes. Record the absorbance at 600 nm, 25 C, to register a steady rate and then add to both tubes freshly prepared NADH 10 mM. Mix lightly and record the absorbance rate.
Since the extinction coefficient for DCIP is 19.1 M/cm, the specific activity of Complex I is given by: Activity ðnmol min mLÞ ¼ ΔAbs½S ΔAbs½Blk Vt 1000 = EC Δtime mg C½S ,
Determination of Oxidative Phosphorylation Complexes Activities
25
where ΔAbs[S] is the change in sample absorbance, ΔAbs[Blk] is the change in rotenone tube absorbance, Vt is the final cuvette volume in microliters, mg is the mitochondrial protein content (for this example, it is 0.03), EC is the extinction coefficient (in this example, 19.1), C[S] is the sample concentration, and Δtime is the elapsed time between stable recordings of absorbance. 3.4.2 Complex II (Succinate:Ubiquinone Oxidoreductase)
Complex II is a particular protein complex, which has four major differences from the other complexes of the respiratory chain. Its subunits are coded exclusively in the nuclear genome, is not a transmembrane protein (i.e., it is only present in the matrix side of the inner mitochondrial membrane and in its interior), its activity does not result in direct proton ejection (although the electrons it collects from oxidizing succinate can lead to protonic ejection at the level of Complexes III and IV), and, as its name implies, it is also part of the Krebs cycle or citric acid cycle. To evaluate the activity of Complex II, we describe a method first published by [6]. Contrary to Complex I (and III and V, as seen later), the activities of Complexes II and IV are polarographically evaluated, with resource to a Clark-type oxygen electrode, connected to a digital or analogic register. In this method, Complex II accepts electrons from succinate, and since mitochondria are burst due to the freeze-thaw cycle, it does not donate electrons to ubiquinone but rather to the supplied acceptor, phenazine methosulfate (PMS). PMS will then, in turn, donate the electrons directly to molecular oxygen, causing its conversion to water. 1. Prepare respiration buffer as above (as described in Subheading 2.1). 2. Depending on the volume of the oxygen electrode chamber, use at least 1 mL of respiration buffer supplemented with: (a) Succinate 5 mM. (b) Rotenone 2 μM (Complex I inhibitor). (c) Antimycin A 0.1 μg/mL of final volume (Complex III inhibitor). (d) KCN 1 mM (Complex IV inhibitor). (e) Triton X-100 0.3 mg/mL of final volume. (f) 0.3 mg freeze-thawed mitochondrial protein. 3. The reaction is initiated by the addition of PMS 1 mM (see Note 17). 4. Complex II activity is measured by calculating the slope of oxygen decrease registered by the electrode, in its initial phase, as demonstrated by Fig. 2, and is typically presented in nAtoms O/min/mg protein (see Note 18).
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Joa˜o S. Teodoro et al.
Fig. 2 Typical polarographic register for the evaluation of Complexes II and IV activities. The red line denotes the oxygen presence in the chamber. The arrow marks the addition of the reaction initiator (PMS for II and ascorbate + TMPD for IV). The dashed line is then drawn to calculate the slope of oxygen disappearance 3.4.3 Complex III (Coenzyme Q: Cytochrome c Oxidoreductase)
Complex III of the mitochondrial respiratory chain is responsible for transferring electrons from fully reduced ubiquinone (known as ubiquinol) to the soluble protein cytochrome c, with a consequent protonic ejection. As mentioned before, the activity of Complex III can be spectrophotometrically evaluated using the method first described by [7] and further explored by [1]. In this method, similarly as before, one follows the increase in absorbance at 550 nm caused by the formation of reduced cytochrome c. l
Prepare respiration buffer as above (described in Subheading 2.1).
l
Use a solution of decylubiquinone to freshly prepare decylubiquinol (see Note 19).
l
To 1 mL of respiration buffer, add 0.3 mg of freeze-thawed mitochondrial protein, decylubiquinol 80 mM, KCN 240 μM, rotenone 4 μM, and ATP 200 μM. Prepare two tubes, one with and the other without antimycin A 0.1 μg/mL of final volume (see Note 20).
l
Start the reaction by adding oxidized cytochrome c 40 μM and then measure the rate of absorbance.
Since the extinction coefficient for reduced cytochrome c is 0.021 M/cm, the specific activity of Complex III is given by: Activity ðnmol min mLÞ ¼ ΔAbs½S ΔAbs½Blk Vt 1000 = EC Δtime mg C½S , where ΔAbs[S] is the change in sample absorbance, ΔAbs[Blk] is the change in rotenone tube absorbance, Vt is the final cuvette volume in microliters, mg is the mitochondrial protein content (for this
Determination of Oxidative Phosphorylation Complexes Activities
27
example, it is 0.03), EC is the extinction coefficient (in this example, 0.021), C[S] is the sample concentration, and Δtime is the elapsed time between stable recordings of absorbance. 3.4.4 Complex IV (Cytochrome c Reductase)
Complex IV is the last protein complex of the electronic transport chain. It receives electrons from cytochrome c and supplies them to molecular oxygen, generating water. This electric transport, as with Complexes I and III, is accompanied by the protonic ejection from the matrix to the intermembrane space. As with Complex II, Complex IV’s activity can be polarographically evaluated using the method first described by [8]. This method also implies oxygen consumption, but since that is a natural reaction for Complex IV, no artificial electron acceptor is required. The reaction is also supplemented with a mix of ascorbate + TMPD (see Note 21). 1. Prepare respiration buffer as above (described in Subheading 2.1). 2. Depending on the volume of the oxygen electrode chamber, use at least 1 mL of respiration buffer supplemented with: (a) Rotenone 2 μM (Complex I inhibitor). (b) Oxidized cytochrome c 10 μM. (c) Triton X-100 0.3 mg/mL of final volume. (d) 0.3 mg freeze-thawed mitochondrial protein. 3. Start the reaction by adding ascorbate 5 mM + TMPD 0.25 mM (see Note 22). 4. As with Complex II, activity should be evaluated by calculating the slope of oxygen decrease registered by the electrode, in its initial phase, as demonstrated by Fig. 2, and is typically presented in nAtoms O/min/mg protein.
3.4.5 ATPase Activity of Mitochondrial ATPSynthase
Mitochondrial ATPSynthase in the enzymatic complex utilizes the electrochemical gradient generated by the respiratory chain to drive ADP phosphorylation. The potential energy stored in such gradient is utilized by ATPSynthase to drive the generation of a chemical bond between ADP and a free phosphate group. The protons are then returned to the matrix, where they can be used, either by the respiratory chain or other enzymes inside mitochondria. It has been shown that, when the membrane potential is sufficiently reduced or completely destroyed, ATPSynthase can revert its activity, that is, it has been shown that it will utilize ATP to pump protons out of the mitochondrial matrix [9]. This property can be explored to assess the activity of this enzymatic complex. l
Produce ATPase reaction buffer as above (described in Subheading 2.1).
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Table 2 ATPase reaction tubes Tube
Reaction buffer (mL)
Mitochondria (mg)
Oligomycin (μg)
X1
2
0.25
0
X10
2
0.25
0.25
Table 3 ATPase reaction standard curve Tube
KH2PO4 0.5 mM (mL)
H2O (mL)
nmol of Pi in tube
0
0
3
0
1
0.25
2.75
125
2
0.5
2.5
250
3
1
2
500
l
Produce trichloroacetic acid (TCA) 40% (v/v), KH2PO4 0.5 mM, and Molybdate reagent 10% in H2SO4 10 N (as described in Subheading 2.1).
l
Prepare the following tubes, as seen in Table 2, two for each sample (see Note 23).
l
Place the tubes in a water bath at 37 C. Start the reaction in each tube with ATP 2.5 mM.
l
After precisely 10 min, stop each reaction with 1 mL TCA. Mix with an orbital vortex and put the tubes on ice. Collect 1 mL (of the three inside the tube) into a new tube.
l
At this time, prepare a standard curve using Table 3.
l
After preparing all the tubes (samples and standard curve), add 2 mL of Molybdate reagent to each. Vortex and let it react for 3 min on the bench. Read absorbance at 660 nm.
The ATPase activity is calculated as before, for Biuret reaction. Plot the standard curve absorbance against the Table 3 supplied Pi values. Again, an r2 value under 0.995 indicates a nonacceptable curve. Calculate the slope and the intercept. Subtract the intercept to the sample’s absorbance and divide the result by the slope. This value is the nmol of Pi inside the tube. Multiply this value by 4 (to normalize to 1 mg of protein) and then again by 3 (because of the 3 mL inside the first reaction tube, only one was used) and divide the result by 10 (to normalize to 1 min). Do this to all sample tubes and subtract the oligomycin tubes to their respective no-oligomycin tube. The result is the specific activity of the mitochondrial ATPase. In sum,
Determination of Oxidative Phosphorylation Complexes Activities
29
Mitochondrial ATPase activity ðnmol Pi=mg protein= min Þ ¼ Abs½S Ins =Slp 1:2 Abs½O Ins =Slp 1:2 , where Abs[S] is the sample absorbance, Abs[O] is the oligomycin sample absorbance, Ins is the intercept of the standard curve, and Slp is the slope of the standard curve.
4
Notes 1. Do not worry if the sodium potassium tartrate tetrahydrate does not dissolve totally, for the following addition of sodium hydroxide will guarantee its dissolution. 2. It is vital that from the moment the animal is sacrificed to this point to take as less time as possible (roughly 60–90 s should suffice). 3. Notice that this will release a lot of blood from the liver, and as such, the buffer should be replaced in order to remove it (2–3 times for a normal 10 g liver from a 250 g animal should suffice). Take care to not waste liver material when throwing away the old buffer. 4. The pestle should reach the bottom of the homogenizer, which might not be possible at the first stroke; nevertheless, if the tissue was correctly chopped, at the second stroke it should be possible. 5. This centrifugation allows for the sedimentation of nuclei, red cells, broken and intact cells, and other heavier, unwanted components. 6. Waste a small amount of supernatant at the end of the tube to avoid carryover of the pelleted material. 7. To resuspend the mitochondrial pellet, use, for example, a watercolor thin paintbrush. The mitochondrial pellet forms a soft brown layer adhered against the tube wall, with a possible dark red central spot, which can be discarded as it consists of pelleted red blood cell contents. It can also have a superficial mobile layer of pelleted mitochondria, which should be discarded (most of it is easily removed when decanting the buffer) as it is formed by damaged mitochondria. 8. Buffer of 3 or 4 mL should suffice for a typical 10 g liver. 9. This is important to allow the stabilization of the mitochondrial preparation. Use this time to quantify the mitochondrial preparation. A complete quantification should measure not only the protein content, but also the activity of citrate synthase, a known mitochondrial marker. 10. Prepare BSA in water, store at 20 C in aliquots. DOC sodium salt is also prepared in water and stored in a lightprotected glass bottle. Start by the water, wash buffer and
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DOC. When all is ready, add the samples to the respective tubes, making at least duplicates. Only when everything is added, add the Biuret reagent. 11. Start by tube 0 and use it to “zero” the spectrophotometer. The r2 (coefficient of determination) is easily calculated using a standard spreadsheet software and should be between 0.995 and 1 (below 0.995, reject the curve and start again). Using the software again, calculate the intersection of the curve to the yaxis, the curve slope and register the dilution factor (for this example, the dilution factor is 20). 12. The replicates should be roughly similar, that is, no more than 0.05 absorbance units apart—if they are, reject your tubes and start again. 13. As an example, a slope of 0.1240, intersection of 0.0113333, a dilution factor of 20, and an average sample absorbance of 0.5110 indicate that the sample has a protein content of 84.2473 mg/mL. 14. This will cause an immediate shift in the absorbance, creating a rate. It is important to record this rate, for it is recommended that only one cuvette be tackled at a time. 15. As before, this will cause an immediate rate shift, for the same precautions are recommended. 16. The tubes with rotenone serve as a negative control and should not demonstrate any increase in absorbance. 17. Reactions should take place at 25 C, in a light-protected environment, under magnetic stirring, since PMS is a highly light and oxygen-sensitive reagent, for it should be made fresh and used in under a few hours after preparation (or made and kept at 20 C for a few days, protected from the light). A distinct shift in color from yellow to green from PMS indicates the reagents’ decay. 18. A digital oxygen recorder automatically calculates the slope and, as such, indicates the oxygen consumption rate per unit of time. As for analogic measurements, one must know how much oxygen there was inside the chamber dissolved in the buffer. Since pure water, at 1 atm and 25 C, has roughly 2.06 mM of oxygen atoms dissolved within, it is simple to calculate the slope in terms of time. 19. To the previously described decylubiquinone solution, add a few crystals of lithium borohydride and mix thoroughly by pipetting until the solution is clear. If excess crystals are added, a few drops of concentrated HCl should get rid of them. When HCl is added and no bubbling is visible, the solution is ready. This will cause the final pH to be between 2 and 3.
Determination of Oxidative Phosphorylation Complexes Activities
31
20. Antimycin A is a specific Complex III inhibitor, and this will be, as before, the negative control tube. 21. Ascorbate helps maintain TMPD in the reduced state, which supplies electrons to cytochrome c. 22. Reactions should take place at 25 C, in a light-protected environment, under magnetic stirring. Ascorbate + TMPD can be added simultaneously but should be kept at 20 C and light-protected, preferably freshly made prior to being used. 23. Use only glass test tubes that have been submerged in concentrated HCl (2 M) for at least 24 h. Wash the tubes after the acid bath with ultrapure water. References 1. Barrientos A, Fontanesi F, Diaz F (2009) Evaluation of the mitochondrial respiratory chain and oxidative phosphorylation system using polarography and spectrophotometric enzyme assays. Curr Protoc Hum Genet Chapter 19:Unit19.3 2. Gornall AG, Bardawill CJ, David MM (1949) Determination of serum proteins by means of the biuret reaction. J Biol Chem 177 (2):751–766 3. Janssen R, Nijtmans LG, van den Heuvel LP, Smeitink JA (2006) Mitochondrial complex I: structure, function and pathology. J Inherit Metab Dis 29(4):499–515 4. Janssen AJM, Trijbels FJM, Sengers RCA, Smeitink JAM, van den Heuvel LP, Wintjes LTM, Stoltenborg-Hogenkamp BJM, Rodenburg RJT (2007) Spectrophotometric assay for complex I of the respiratory chain in tissue samples and cultured fibroblasts. Clin Chem 53 (4):729–734
5. Fischer JC, Ruitenbeek W, Trijbels JM, Veerkamp JH, Stadhouders AM, Sengers RC, Janssen AJ (1986) Estimation of NADH oxidation in human skeletal muscle mitochondria. Clin Chim Acta 155(3):263–273 6. Singer TP (1974) Determination of the activity of succinate, NADH, choline, and alphaglycerophosphate dehydrogenases. Methods Biochem Anal 22:123–175 7. Tisdale HD (1967) Preparation and properties of succinic-cytochrome c reductase (complex II-III). Methods Enzymol 10:213–215 8. Brautigan DL, Ferguson-Miller S, Margoliash E (1978) Mitochondrial cytochrome c: preparation and activity of native and chemically modified cytochromes c. Meth Enzymol 53:128–164 9. Jonckheere AI, Smeitink JAM, Rodenburg RJT (2012) Mitochondrial ATP synthase: architecture, function and pathology. J Inherit Metab Dis 35(2):211–225
Chapter 3 BN-PAGE-Based Approach to Study Thyroid Hormones and Mitochondrial Function Elena Silvestri, Assunta Lombardi, Federica Cioffi, and Fernando Goglia Abstract In recent years, a number of advancements have been made in the study of entire mitochondrial proteomes in both physiological and pathological conditions. Naturally occurring iodothyronines (i.e., T3 and T2) greatly influence mitochondrial oxidative capacity, directly or indirectly affecting the structure and function of the respiratory chain components. Blue native PAGE (BN-PAGE) can be used to isolate enzymatically active oxidative phosphorylation (OXPHOS) complexes in one step, allowing the clinical diagnosis of mitochondrial metabolism by monitoring OXPHOS catalytic and/or structural features. Protocols for isolating mammalian liver mitochondria and subsequent one-dimensional (1D) BN-PAGE will be described in relation to the impact of thyroid hormones on mitochondrial bioenergetics. Key words Thyroid hormone, Iodothyronine, Mitochondrion, Respiratory chain, BN-PAGE
1
Introduction Mitochondria are “hybrid” and dynamic organelles resulting from the coordinated expression of both the nuclear and their own genome [1, 2], critically relevant for energy homeostasis, metabolism, regulation of apoptosis, and proper cell viability. Physiologically, mitochondrial functions are ensured and orchestrated by metabolic (i.e., hormones), environmental, and developmental signals, allowing tissues to adjust their energy production according to changing demands. Understanding how mitochondria follow these adjustments would provide invaluable information and give insight into both mitochondrial functions and mitochondria-associated diseases. Very recent studies, over the last decade, specifically clarified how mitochondria are not only highly mobile, but also shape-changing organelles in which macro and ultrastructural modifications directly regulate bioenergetic functions [3– 5]. Importantly, mitochondria are key subcellular targets for
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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thyroid hormones (THs; thyroxine (T4) and 3,30 ,5-triiodo-L-thyronine (T3)) and iodothyronines (i.e., 3,5-diiodo-L-thyronine (T2)) that regulate energy and substrate utilization, which are closely dependent on their effects on mitochondrial functions [6, 7]. Indeed, extensive changes occur in the mitochondrial compartment in response either to THs or to physiological/pathological states, involving changes in the activity of the thyroid gland [7, 8]. The impact of THs on mitochondrial function is particularly evident in metabolically active tissues, including skeletal muscle, heart, kidney, and liver. The liver, in particular, has a central role in the potent hypolipidemic effect of both T3 and T2 [9–11], which exert a strong inhibitory effect on the development of steatosis [9, 12, 13]. This antisteatotic effect implies a facilitation of fatty acid transport, an induction of fatty acid oxidation, and a reduction of the severity of liver injury determined by the serum levels of transaminases, suggesting a common mechanism of mitochondrial metabolism modulation [9, 12, 13]. To study the response of mitochondria to THs under physiological, pathological, and pharmacological conditions, an integrated multidisciplinary approach is required [14]. Indeed, the high level of compartmentalization in mitochondria and the existence of multipolypeptide complexes that contain hydrophobic proteins in close contact with the membrane lipids, peripheral proteins, and nonprotein cofactors imply that a deep structural/ functional study of the mitoproteome requires an appropriate combination of different tools to compensate for the limits imposed by each individual technique [14]. During the last decade, proteomics has been increasingly used to identify and quantify mitochondrial proteins related to cellular perturbations, enforcing data from metabolites and gene sequences both in physiological and in pathological situations [15]. To study the effects of iodothyronine administration on total tissue and subcellular compartments in metabolically active tissues, we performed high-resolution differential proteomic analyses combining two-dimensional gel electrophoresis (2D-E) and subsequent matrix-assisted laser desorption/ ionization time-of-flight mass spectrometry (MALDI-TOF MS) and nanoliquid chromatography-electrospray ionization-linear ion trap (LC-ESI-LIT)-MS/MS techniques [10, 16–18]. To gain deeper insights into both the mitochondrial response mechanisms to iodothyronines and the resulting proteome alterations, we employed one-dimensional (1D) blue native PAGE [1D BNPAGE] to examine the abundance of mitochondrial OXPHOS complexes as well as their supramolecular organization and activity [10, 19–21]. In particular, BN-PAGE was used to separate mitochondrial proteins and complexes in the mass range of 10 kDa– 10 MDa [21–24]. Briefly, BN-PAGE is a discontinuous microscale electrophoretic technique to isolate microgram amounts of
Study of Liver Mitochondrial OXPHOS by (1D) BN-PAGE
a
35
b KDa 6% 1000
violet
Complex V
700
white
Complex III dim
BN gradient gel
Complex I
490
200
Complex IV
130
Complex II
brown pink
13%
Fig. 1 Representative BN-PAGE separation of dodecyl maltoside-solubilized mitochondrial complexes. (a) Coomassie blue stained BN-PAGE 6–13% gradient gel of crude rat liver mitochondria. Bands characteristic of individual OXPHOS complexes are recognizable. The native mass range is also reported. 15 μg of mitochondrial protein extract were loaded. (b) Histochemical staining of in-gel activity of individual OXPHOS complexes. The color of the specific band staining is reported
membrane protein complexes from biological membranes for use several uses, such as the clinical diagnostics of mitochondrial disorders, identification of protein-protein interactions, in-gel activity, protein import, etc. ([23], and references within). For BN-PAGE, membranes are solubilized by nonionic (neutral), nondenaturing detergents, selected based on the detergent stability of the complexes of interest (e.g., digitonin is the mildest detergent, dodecyl maltoside has stronger delipidating properties, and Triton X-100 shows intermediate behavior). After solubilization, Coomassie Blue G-250 is added, which binds to the surface of the proteins and converts them into water-soluble molecules. This allows the negatively charged complexes or supercomplexes to be separated according to molecular mass and detected as blue bands in the BN gels (Figs. 1 and 2). Supramolecular assemblies retained from the 1D BN-PAGE can be dissociated into individual complexes by applying an orthogonal-modified BN-PAGE for a second native dimension, allowing the identification of interacting partners as well as their stoichiometric ratio [25]. Because 1D BN-PAGE separates intact OXPHOS complexes/supercomplexes, subsequent denaturing electrophoresis can resolve the individual subunits of the respective complexes. Indeed, OXPHOS assembly profiles can be obtained by
Elena Silvestri et al.
KDa
SC1 (containing complex I activity)
6% 2000
SC2 (containing complex I activity) SC3 (containing complex I and IV activities) SC4 (containing complex IV activity)
1500
BN gradient gel
36
1200 890
380
SC5 (containing complex IV activity)
200 13%
Fig. 2 Representative BN-PAGE separation of digitonin-solubilized mitochondrial proteins. Coomassie blue stained BN-PAGE 6–13% gradient gel of crude rat liver mitochondria. Bands of OXPHOS supercomplexes (SC) are recognizable. The native mass range is also reported. 15 μg of mitochondrial protein extract were loaded per lane. Arrows and numbers indicate bands of OXPHOS SCs in which Complex I and Complex IV activities were detected by histochemical staining
two-dimensional blue native/SDS gel electrophoresis, which provides additional information on the role of specific proteins in the biogenesis of the OXPHOS system [23, 26]. Overall, BN-PAGE is a robust, manageable, reproducible, and cost- and time-efficient method, and it has the important advantage of being compatible with in-gel activity staining procedures, specifically measuring the OXPHOS enzymes (whereas most spectrophotometric assays also detect other cellular activities) [27–30] (Fig. 1). A number of previous studies support how BN-PAGE, combined with histochemical staining, can provide valuable information for the clinical diagnosis of OXPHOS states by monitoring their catalytic and/or structural features [31–33]. Of note, the measurement of the specific activities in reactive bands has to be considered semiquantitative and strongly affected by the in-gel milieu [24, 34, 35]. The preparation of mitochondria from tissues such as liver, as well as the subsequent processing for BN-PAGE, can be problematic and actually differ somewhat from the protocols for muscle and brain tissue. Here, we will describe the protocols used in our laboratory to isolate the mitochondria from mammalian liver and to study OXPHOS complexes by 1D BN-PAGE in terms of assembly and individual in-gel activity.
Study of Liver Mitochondrial OXPHOS by (1D) BN-PAGE
2
37
Materials
2.1 Materials and Equipment
1. Prepare all solutions using ultrapure (double-distilled (DD)) water and analytical grade reagents (we did not add sodium azide to the reagents) and store all reagents at 4–8 C (unless indicated otherwise). Diligently follow all waste disposal regulations. 2. The following pieces of equipment are required: a commercially available vertical electrophoresis apparatus (optional, (mini)gel multicasting chamber), a power supply (600 V, 500 mA), a motor-driven tightly fitting glass–Teflon homogenizer, a gradient mixer for use with a magnetic stirrer (optional peristaltic pump for casting acrylamide gradient gels), refrigerated centrifuges, and ultracentrifuges. We most often use minigel systems in our laboratory, which allow sufficiently good separation of the OXPHOS complexes and have the advantage of requiring fewer reagents and less sample material. This is important for downstream applications such as in-gel activity assays, which necessitate expensive chemicals.
2.2 Mitochondria Isolation and BN-PAGE Sample Preparation
1. Phenylmethylsulfonyl fluoride (PMSF) stock solution: 0.5 M in dimethylsulfoxide (DMSO). 2. Sucrose buffer: 440 mM sucrose, 20 mM MOPS, 1 mM EDTA, pH 7.2, and 0.2 mM PMSF (to be added shortly before use). 3. NaCl solution: 500 mM NaCl, 10 mM Na+/MOPS, pH 7.2. 4. Aminocaproic acid/Bis-Tris HCl solution: 1 M aminocaproic acid, 50 mM Bis-Tris HCl, pH 7. 5. Triton X-100 solution: 10%. 6. Dodecyl maltoside solution: 10% dodecyl maltoside (w/v) in water; store 1 ml aliquots at 20 C. 7. Digitonin solution: 20% digitonin (w/v) in water; store 0.1–1 ml aliquots at 20 C. Heating (>70 C) can be required for some lots of digitonin but avoid repeated freezing/thawing, which can lead to the insolubility of digitonin at temperatures 620 nm long pass (FL-3/far red). Sperm-specific events of 15,000–50,000 per condition should be recorded and further analyzed with an adequate software. Nonsperm events must be gated out, as previously described [28]. 3.1.3 MMP Evaluation
1. Incubate 2.5–5 106 of washed spermatozoa/mL in sperm medium with the fluorescent probe at 37 C in the dark (see Note 2). For MMP assessment, an appropriate control should be used. Check Table 2 for probes and control concentrations and times of incubation; 2. After incubation, centrifuge the suspension at 300–500 g for 5 min; 3. Discard the supernatant and resuspend the cells in an appropriate volume of sperm medium (e.g., 100μL for fluorescence microscopy and 300–500μL for flow cytometry). 4. For fluorescence microscopy analysis: (a) Place 10μL of sperm suspension on a glass slide under a coverslip and observe the fluorescence with 546/12 filter for TMRM and filters 546/12 and 486/20 for JC-1; (b) Count 200 sperm cells in different areas of the slide to determine the percentage of labeled cells. Check Fig. 2 for representative images of human sperm labeled with MMP fluorescent probes. 5. For flow cytometry experiments: (a) For data acquisition, a cytometer is required with an argon laser that performs with an excitation wavelength of 488 nm coupled with the following emission filters: 530/30 band pass (FL-1 channel/green), 585/42 band
Table 2 Fluorescent probes for MMP assessment Fluorescent probe
Concentration
Time of incubation (min)
Control
Oocytes
JC-1
1.8μM
40
CCCP (50μM)
Sperm
JC-1 TMRM
2μM 300 ηM
20 25
CCCP (50μM) CCCP (50μM)
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Renata S. Tavares et al.
pass (FL-2 channel/red), and >620 nm long pass (FL-3/ far red). Sperm-specific events of 15,000–50,000 per condition should be recorded and further analyzed with an adequate software. Nonsperm events must be gated out, as previously described [28]. 3.2
Oocytes
3.2.1 Oocytes Preparation
1. Place the cumulus-oocyte complexes (COC) in the first well of a 4-well plate previously filled with PBS; 2. Denudate COCs by successive pipetting the cells up and down in PBS; 3. Repeat step 2 sequentially in the other three wells in order to wash the oocytes and ensure the complete removal of all cumulus cell layers.
3.2.2 Mitochondrial H2O2 Levels
1. Incubate the denudated oocytes in a new plate with oocyte medium containing MitoPY1 at 37 C in maximum humidity. For MitoPY1 assessment, an appropriate control should be used. Check Table 1 for probe and control concentration and time of incubation; 2. Wash the oocytes, by successive pipetting the cells up and down in PBS, to ensure the removal of probe excess; 3. Place the oocytes on glass slides under a coverslip and examine in a fluorescence microscope; 4. Observe the fluorescence with a 486/20 filter.
3.2.3 MMP Evaluation
1. Incubate the denudated oocytes in a new plate with oocyte medium containing JC-1 at 37 C in maximum humidity. For MMP assessment, an appropriate control should be used. Check Table 2 for probe and control concentration and time of incubation; 2. Wash the oocytes, by successive pipetting the cells up and down in PBS, to ensure the removal of probe excess; 3. Place the oocytes on glass slides under a coverslip and examine in a fluorescence microscope; 4. Observe the fluorescence with a 486/20 filter.
3.2.4 Mitochondrial Mass/Distribution/ Aggregation
1. Incubate the denudated oocytes in oocyte medium containing 30 nM of MitoTracker™ Green for 30 min at 37 C in maximum humidity (also see Note 1); 2. Wash the oocytes, by successive pipetting the cells up and down in PBS, to ensure the removal of probe excess; 3. Place the oocytes on glass slides under a coverslip and examine under a fluorescence microscope; 4. Observe the fluorescence with a 486/20 filter.
Mitochondrial Probes for Gametes
4
67
Notes 1. MitoTracker™ Green is meant to evaluate total mitochondrial mass, and it accumulates inside the mitochondria regardless of MMP; therefore, we see no need for a control. 2. For fluorescence microscopy, when incubating the cells with the probes, add Hoechst 33342 (5μg/mL) in the final 5 min (for sperm cells) or 15 min (for oocytes) of incubation to facilitate cell observation under the microscope. Observe the blue fluorescence with a 365/12 filter. 3. In Fig. 1, the nuclear staining observed when using MitoSOX™ Red in sperm cells is expected, since, although this is a probe that detects mitochondrial superoxide anion, the reacted probe binds to DNA and can, therefore, stain the sperm nucleus. In fact, this type of labeling has been previously reported and is commonly accepted in these cells [4].
References 1. Ramalho-Santos J, Varum S, Amaral S, Mota PC, Sousa AP, Amaral A (2009) Mitochondrial functionality in reproduction: from gonads and gametes to embryos and embryonic stem cells. Hum Reprod Update 15:553–572 2. Gosalvez J, Tvrda E, Agarwal A (2017) Free radical and superoxide reactivity detection in semen quality assessment: past, present, and future. J Assist Reprod Genet 34:697–707 3. Sasaki H, Hamatani T, Kamijo S, Iwai M, Kobanawa M, Ogawa S, Miyado K, Tanaka M (2019) Impact of oxidative stress on age-associated decline in oocyte developmental competence. Front Endocrinol (Lausanne) 10:811 4. Koppers AJ, De Iuliis GN, Finnie JM, McLaughlin EA, Aitken RJ (2008) Significance of mitochondrial reactive oxygen species in the generation of oxidative stress in spermatozoa. J Clin Endocrinol Metab 93:3199–3207 5. De Jonge CJ, Barratt CLR (2006) The sperm cell: production, maturation, fertilization, regeneration. Cambridge University Press, Cambridge 6. Agarwal A, Virk G, Ong C, du Plessis SS (2014) Effect of oxidative stress on male reproduction. World J Mens Health 32:1 7. du Plessis SS, McAllister DA, Luu A, Savia J, Agarwal A, Lampiao F (2010) Effects of H(2) O(2) exposure on human sperm motility parameters, reactive oxygen species levels and nitric oxide levels. Andrologia 42:206–210
8. Goud PT, Goud AP, Diamond MP, Gonik B, Abu-Soud HM (2008) Nitric oxide extends the oocyte temporal window for optimal fertilization. Free Radic Biol Med 45:453–459 9. Amaral A, Ramalho-Santos J (2010) Assessment of mitochondrial potential: implications for the correct monitoring of human sperm function. Int J Androl 33:180–186 10. Van Blerkom J (2002) Domains of highpolarized and low-polarized mitochondria may occur in mouse and human oocytes and early embryos. Hum Reprod 17:393–406 11. Sun QY, Wu GM, Lai L, Park KW, Cabot R, Cheong HT, Day BN, Prather RS, Schatten H (2001) Translocation of active mitochondria during pig oocyte maturation, fertilization and early embryo development in vitro. Reproduction 122:155–163 12. Dalton CM, Carroll J (2013) Biased inheritance of mitochondria during asymmetric cell division in the mouse oocyte. J Cell Sci 126:2955–2964 13. Purdey MS, Connaughton HS, Whiting S, Schartner EP, Monro TM, Thompson JG, Aitken RJ, Abell AD (2015) Boronate probes for the detection of hydrogen peroxide release from human spermatozoa. Free Radic Biol Med 81:69–76 14. Portela JMD, Tavares RS, Mota PC, RamalhoSantos J, Amaral S (2015) High glucose concentrations per se do not adversely affect human sperm function in vitro. Reproduction 150:77–84
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15. Sousa MI, Amaral S, Tavares RS, Paiva C, Ramalho-Santos J (2014) Concentrationdependent sildenafil citrate (Viagra) effects on ROS production, energy status, and human sperm function. Syst Biol Reprod Med 60:72–79 16. Amaral S, Redmann K, Sanchez V, Mallidis C, Ramalho-Santos J, Schlatt S (2013) UVB irradiation as a tool to assess ROS-induced damage in human spermatozoa. Andrology 1:707–714 17. Aitken RJ, Smith TB, Lord T, Kuczera L, Koppers AJ, Naumovski N, Connaughton H, Baker MA, De Iuliis GN (2013) On methods for the detection of reactive oxygen species generation by human spermatozoa: analysis of the cellular responses to catechol oestrogen, lipid aldehyde, menadione and arachidonic acid. Andrology 1:192–205 18. Mukhopadhyay P, Rajesh M, Hasko G, Hawkins BJ, Madesh M, Pacher P (2007) Simultaneous detection of apoptosis and mitochondrial superoxide production in live cells by flow cytometry and confocal microscopy. Nat Protoc 2:2295–2301 19. Dickinson BC, Lin VS, Chang CJ (2013) Preparation and use of MitoPY1 for imaging hydrogen peroxide in mitochondria of live cells. Nat Protoc 8:1249–1259 20. Escada-Rebelo S, Mora F, Sousa A, AlmeidaSantos T, Paiva A, Ramalho-Santos J (2020) Fluorescent probes for the detection of reactive oxygen species in human spermatozoa. Asian J Androl 22(5):465–471. https://doi.org/10. 4103/aja.aja_132_19 21. Paoli D, Gallo M, Rizzo F, Baldi E, Francavilla S, Lenzi A, Lombardo F, Gandini L (2011) Mitochondrial membrane potential profile and its correlation with increasing sperm motility. Fertil Steril 95:2315–2319 22. Troiano L, Granata ARM, Cossarizza A, Kalashnikova G, Bianchi R, Pini G, Tropea F,
Carani C, Franceschi C (1998) Mitochondrial membrane potential and DNA stainability in human sperm cells: a flow cytometry analysis with implications for male infertility. Exp Cell Res 241:384–393 23. Uribe P, Villegas JV, Boguen R, Treulen F, Sa´nchez R, Mallmann P, Isachenko V, Rahimi G, Isachenko E (2017) Use of the fluorescent dye tetramethylrhodamine methyl ester perchlorate for mitochondrial membrane potential assessment in human spermatozoa. Andrologia 49:e12753 24. Baptista M, Publicover SJ, Ramalho-Santos J (2013) In vitro effects of cationic compounds on functional human sperm parameters. Fertil Steril 99:705–712 25. Tavares RS, Amaral S, Paiva C, Baptista M, Ramalho-Santos J (2015) In vitro exposure to the organochlorine p,p’-DDE affects functional human sperm parameters. Chemosphere 120:443–446 26. Liu S, Li Y, Gao X, Yan JH, Chen ZJ (2010) Changes in the distribution of mitochondria before and after in vitro maturation of human oocytes and the effect of in vitro maturation on mitochondria distribution. Fertil Steril 93:1550–1555 27. Ramalho-Santos J, Amaral A, Sousa AP, Rodrigues AS, Martins L, Baptista M, Mota PC, Tavares R, Amaral S, Gamboa S (2007) Probing the structure and function of mammalian sperm using optical and fluorescence microscopy. Modern Res Educ Topics Microsc 1:394–402 28. Sousa AP, Amaral A, Baptista M, Tavares R, Caballero Campo P, Caballero Peregrin P, Freitas A, Paiva A, Almeida-Santos T, Ramalho-Santos J (2011) Not all sperm are equal: functional mitochondria characterize a subpopulation of human sperm with better fertilization potential. PLoS One 6:e18112
Chapter 6 Update on the Histoenzymatic Methods for Visualization of the Activity of Individual Mitochondrial Respiratory Chain Complexes in the Human Frozen Sections Mariusz R. Wieckowski, Maciej Pronicki, and Agnieszka Karkucinska-Wieckowska Abstract Investigation of mitochondrial metabolism perturbations and successful diagnosis of patients with mitochondrial abnormalities often requires assessment of human samples like muscle or liver biopsy as well as autopsy material. Immunohistochemical and histochemical examination is an important technique to investigate mitochondrial dysfunction that combined with spectrophotometric and Blue Native electrophoresis techniques can be an important tool to provide diagnosis of mitochondrial disorders. In this chapter, we focus on technical description of the methods that are suitable to detect the activity of complex I, II, and IV of mitochondrial respiratory chain in frozen sections of brain, heart, muscle, and liver biopsies/autopsy. The protocols provided can be useful not only for general assessment of mitochondrial activity in studied material, but they are also successfully used in the diagnostic procedures in case of suspicion of mitochondrial disorders. In the age of high-performance NGS sequencing, these methods can be used to confirm whether mutations are pathogenic by proving their impact on the activity of individual respiratory chain complexes. Key words Histoenzymatic methods, Frozen sections, Mitochondrial respiratory chain complexes
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Introduction As the center of oxidative metabolism, mitochondria are considered as a cellular powerhouse. They produce the majority of cellular ATP, but also, mitochondria are involved in several other critical metabolic processes [1]. For this reason, and many others, defects in mitochondrial function are often connected with pathological states and can lead to various diseases [2–5]. Special attention should be devoted to the alterations in the mitochondrial respiratory chain. Localization and measurement of changes in the activity of respiratory chain complexes can provide important information about the disease etiology. A wide spectrum of methods enables
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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studying the activity of respiratory chain complexes in isolated mitochondria, intact cells, as well as in tissues samples [6– 8]. Under special experimental conditions, the activity of individual mitochondrial respiratory chain complex can be measured separately using spectrophotometric methods based on electron donor/acceptor enzymatic-coupled reactions [9]. Blue Native electrophoresis, followed by in-gel activity assay, can also be successfully applied either for detection of respiratory chain complex deficiency or for visualization deficiencies of the individual respiratory chain complex activity [10, 11]. Histoenzymatic methods presented in our chapter are dedicated for visualization of alterations in the activity of individual mitochondrial respiratory chain complexes in the frozen sections from patients for diagnostic purpose [12– 15]. Here, we present examples of histoenzymatic analysis of complex I, II, and IV activity in frozen sections of brain, heart, muscle, and liver biopsies/autopsy (see Note 1).
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Materials
2.1 Assessment of Cytochrome c Oxidase (COX; Complex IV) Activity in the Human Frozen Sections 2.1.1 Solutions and Chemicals
1. Catalase stock: 20μg/ml (see Note 2). 2. Formal-calcium (fixer): 15.8 g of anhydrous calcium chloride dissolved in 60 ml of deionized water and supplemented with 40 ml of 40% formaldehyde (see Note 3). 3. 0.1 M phosphate buffer, pH 7.4 (see Note 4): Directly before use, 4.05 ml of Na2HPO4 solution (1.42 g of Na2HPO4 dissolved in 50 ml of deionized water) should be mixed with 0.95 ml of NaH2PO4 solution (1.2 g NaH2PO4 dissolved in 50 ml of deionized water) and supplemented with 5 ml of deionized water. 4. Incubation solution (to be prepared directly before use): 5 mg of DAB dissolved in 9 ml of 0.1 M phosphate buffer, pH 7.4. Next reagents should be added in the following order: 10 mg cytochrome c (see Note 5), 1 ml of catalase stock, and 750 mg sucrose. Solution should be mixed well without doing foam. 5. Formaldehyde solution for molecular biology, 36.5–38%. 6. Xylene (mixed isomers). 7. Ethanol 96%. 8. Ethanol anhydrous 99.6%. 9. DePeX mountant for histology.
2.1.2 Equipment and Accessories
1. Cryostat (Cryotome FSE). 2. Incubator with the ability to achieve below-ambient temperature.
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3. Olympus: BX53 with XC50 digital camera and CellSense Dimension image-capture software or Hamamatsu NanoZoomer 2.0 RS scanner. 4. Humidity chamber for immunohistochemical staining. 5. Coplin staining jar. 6. Coated microscope slide Super Frost Plus. 7. Cover glasses. 8. DakoPen. 2.2 Assessment of Succinate Dehydrogenase (SDH; Complex II) Activity in the Human Frozen Sections 2.2.1 Solutions and Chemicals
1. Formal-calcium (fixer): 15.8 g of anhydrous calcium chloride dissolved in 60 ml of deionized water and supplemented with 40 ml of 40% formaldehyde (see Note 3). 2. 0.1 M phosphate buffer, pH 7.6 (see Note 4): Use 88 ml of Na2HPO4 solution (1.42 g of Na2HPO4 dissolved in 100 ml of deionized water) should be mixed with 12 ml of KH2PO4 solution (1.36 g KH2PO4 dissolved in 100 ml of deionized water). 3. NBT solution: 100 mg of nitrotetrazolium blue chloride (Sigma N6876), 6.5 mg KCN, and 185 mg EDTA dissolved in 100 ml of 0.1 M phosphate buffer, pH 7.6 (see Note 6). 4. Succinate stock: 2.7 g of sodium succinate dibasic hexahydrate dissolved in 20 ml of deionized water (see Note 7). 5. Incubation solution (prepared directly before use): 0.2 ml of succinate stock mixed with 2 ml of NBT solution and 1 mg of phenazine methosulphate (see Note 8). 6. Formaldehyde solution for molecular biology, 36.5–38%. 7. Xylene (mixed isomers). 8. Ethanol 96%. 9. Ethanol anhydrous 99.6%. 10. DePeX mountant for histology.
2.2.2 Equipment and Accessories
1. Cryostat (Cryotome FSE). 2. Incubator to achieve stable-ambient temperature. 3. Olympus: BX53 with XC50 digital camera and CellSense Dimension image-capture software or Hamamatsu NanoZoomer 2.0 RS scanner. 4. Humidity chamber for immunohistochemical staining. 5. Coplin staining jar. 6. Coated microscope slide Super Frost Plus. 7. Cover glasses. 8. DakoPen. 9. 0.22μm filter unit.
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2.3 Assessment of NADH Dehydrogenase (Diaphorase; Complex I) Activity in the Human Frozen Sections 2.3.1 Solutions and Chemicals
1. NBT stock: 1 mg of nitrotetrazolium blue chloride (Sigma N6876) in 1 ml of deionized water (see Note 9). 2. 1 M Tris–HCl buffer, pH 7.4 (see Note 10): 6.1 g of Tris dissolved in 500 ml of deionized water, supplemented with 37 ml of 1 M HCl solution, and adjusted to 1 liter with the use of deionized water. 3. NADH stock (prepared directly before use): 3 mg of β-nicotinamide adenine dinucleotide, reduced, disodium salt (NADH) dissolved in 3 ml of 1 M Tris–HCl buffer. 4. Incubation solution (prepared directly before use): 3 ml of NADH stock mixed with 3 ml of NBT stock solution. 5. Dako faramount aqueous mounting medium.
2.3.2 Equipment and Accessories
1. Cryostat (Cryotome FSE). 2. Incubator with the ability to achieve below-ambient temperature. 3. Olympus: BX53 with XC50 digital camera and CellSense Dimension image-capture software or Hamamatsu NanoZoomer 2.0 RS scanner. 4. Humidity chamber for immunohistochemical staining. 5. Coplin staining jar. 6. Coated microscope slide Super Frost Plus. 7. Cover glasses. 8. DakoPen.
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Methods
3.1 Brain, Heart, Liver Autopsy Samples and Muscle Biopsy from Patients
1. Skeletal muscle samples were obtained by open muscle biopsy. Brain, heart, and liver samples were obtained by autopsy. 2. Deep muscle biopsies and brain, heart, and liver autopsy samples in tissue-embedding medium (matrix) and freeze in isopentane cooled by liquid nitrogen. 3. Cut with the use of cryostat from 3 to 5 (per coated microscope slide) 8–10μm sections and subject to histoenzymatic staining for COX, SDH, and NADH diaphorase activity. Sections from muscle have to be obtained from transversely orientated muscle blocks.
3.2 Assessment of Cytochrome c Oxidase (COX; Complex IV) Activity in the Human Frozen Sections
In the place where cytochrome c oxidase (complex IV) has an enzymatic activity, DAB molecule is reduced, and DAB-reduced molecule as a brown precipitate can be observed.
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1. Put unfixed cryostat sections on coated microscope slide. 2. With the use of DakoPen, outline cryostat sections in order to avoid flow of incubation solution from the microscope slide. 3. Put microscope slides into the humidity chamber for immunohistochemical staining. 4. Cover outlined cryostat sections with incubation solution and incubate 1 h at 22 C. 5. Wash three times with distilled water. 6. Fix in formal-calcium solution in Coplin staining jar for 15 min at room temp. 7. Dehydrate sections in Coplin staining jar containing 96% ethanol (2 1 min). 8. Next, dehydrate sections in Coplin staining jar containing 99.6% ethanol (2 1 min). 9. Clear in Coplin staining jar containing xylene (mixed isomers) (3 3 min). 10. Mount in DePeX mountant for histology and cover with cover glass (see Note 11). 11. Analyze with light microscope and record images with different magnifications—brown reaction product at the sites of active cytochrome oxidase should be observed. 12. For future analysis, prepare photographic records from stained sections (Fig. 1).
Fig. 1 Histochemical assessment of cytochrome c oxidase (COX; complex IV) activity in frozen sections of brain, heart, muscle, and liver biopsies/autopsy. Light microscopy, original magnification 400X
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3.3 Assessment of Succinate Dehydrogenase (SDH; Complex II) Activity in the Human Frozen Sections
In the place where SDH (complex II) has an enzymatic activity, NBT molecule is reduced, and NBT-reduced molecule as a purple formazan deposit with NBT can be observed. 1. Put unfixed cryostat sections on coated microscope slide. 2. With the use of DakoPen, outline cryostat sections in order to avoid flow of incubation solution from the microscope slide. 3. Put microscope slides into the humidity chamber for immunohistochemical staining. 4. Cover outlined cryostat sections with incubation solution and incubate 15 min at 37 C. 5. Wash three times with distilled water. 6. Fix in formal-calcium solution in Coplin staining jar for 15 min at room temp. 7. Dehydrate sections in Coplin staining jar containing 96% ethanol (2 1 min). 8. Next, dehydrate sections in Coplin staining jar containing 99.6% ethanol (2 1 min). 9. Clear in Coplin staining jar containing xylene (mixed isomers) (3 3 min). 10. Mount in DePeX mountant for histology and cover with cover glass (see Note 11). 11. Analyze with light microscope and record images with different magnifications—bluish-black reaction product at the sites of active succinate dehydrogenase should be observed. 12. For future analysis, prepare photographic records from stained sections (Fig. 2).
3.4 Assessment of NADH Dehydrogenase (Diaphorase; Complex I) Activity in the Human Frozen Sections
In the place where NADH diaphorase (complex I) has an enzymatic activity, NBT molecule is reduced, and NBT-reduced molecule as a purple formazan deposit with NBT can be observed. 1. Put unfixed cryostat sections on coated microscope slide. 2. With the use of DakoPen, outline cryostat sections in order to avoid flow of incubation solution from the microscope slide. 3. Put microscope slides into the humidity chamber for immunohistochemical staining. 4. Cover outlined cryostat sections with incubation solution and incubate 60 min at 37 C. 5. Wash three times with distilled water. 6. Mount in Dako faramount aqueous mounting medium and cover with cover glass (see Note 11).
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Fig. 2 Histochemical assessment of succinate dehydrogenase (SDH; complex II) activity in frozen sections of brain, heart, muscle, and liver biopsies/autopsy. Light microscopy, original magnification 400
7. Analyze with light microscope and record images with different magnifications—bluish-black reaction product at the sites of active NADH diaphorase should be observed. 8. For future analysis, prepare photographic records from stained sections (Fig. 3).
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Notes 1. Ethics: The studies with the use of frozen sections (brain, heart, skeletal muscle, and liver), obtained by muscle biopsy and autopsy examination of brain, heart, and liver, 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/autopsy, or molecular analysis was performed. 2. The stock (approx. 1.2 ml) can be prepared in advance and stored at 20 C for approx. 1 month. 3. Solution can be prepared in advance and stored at room temperature for approx. 1 month in the dark glass bottle. 4. Na2HPO4, NaH2PO4, and KH2PO4 solutions can be prepared in advance and stored at 4 C for approx. 1 month.
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Fig. 3 Histochemical assessment of NADH dehydrogenase (diaphorase; complex I) activity in frozen sections of brain, heart, muscle, and liver biopsies/autopsy. Light microscopy, original magnification 400
5. Use only cytochrome c from horse heart that has been prepared (obtained/purified) without TCA precipitation step. This is very crucial! You can use cyt. c from, for example, SigmaAldrich cat. n. C7752. 6. The NBT solution can be prepared in advance and frozen at 20 C for approx. 1 year in 2.5 ml portions. 7. The succinate stock can be prepared in advance and frozen at 20 C for approx. 1 year in 0.3 ml portions. 8. Mix just before use and keep out of strong light. Solution should be filtered through a 0.22 μm filter unit. 9. The stock of NBT can be prepared in advance and stored at 4 C for approx. 1 month. 10. Solution can be prepared in advance and stored at 4 C for approx. 1 month. 11. Mounted sections can be analyzed immediately or up to 1 year.
Acknowledgments This work was supported by the Internal Projects of The Children’s Memorial Health Institute No. S125/2012 for AKW and MP and by the Polish National Science Centre grant (UMO- 2018/29/B/ NZ1/00589 for MRW).
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References 1. McBride HM, Neuspiel M, Wasiak S (2006) Mitochondria: more than just a powerhouse. Curr Biol 16:R551–R560 2. Lin MT, Beal MF (2006) Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature 443:787–795 3. Maechler P, Wollheim CB (2001) Mitochondrial function in normal and diabetic beta-cells. Nature 414:807–812 4. McKenzie M, Liolitsa D, Hanna MG (2004) Mitochondrial disease: mutations and mechanisms. Neurochem Res 29:589–600 5. Taylor RW, Turnbull DM (2005) Mitochondrial DNA mutations in human disease. Nat Rev Genet 6:389–402 6. Sciacco M, Bonilla E (1996) Cytochemistry and immunocytochemistry of mitochondria in tissue sections. Methods Enzymol 264:509–521 7. Capaldi RA, Murray J, Byrne L, Janes MS, Marusich MF (2004) Immunological approaches to the characterization and diagnosis of mitochondrial disease. Mitochondrion 4:417–426 8. Pronicki M, Szyman´ska-De˛bin´ska T, Karkucin´ska-Wie˛ckowska A, Krysiewicz E, Kaczmarewicz E, Bielecka L, PiekutowskaAbramczuk D, Sykut-Cegielska J, Cukrowska B, Wie˛ckowski MR (2008) Assessment of respiratory chain function in cultured fibroblasts using cytochemistry, immunocytochemistry and SDS-PAGE. Ann Diagn Pediatr Pathol 12:53–61 9. Kramer KA, Oglesbee D, Hartman SJ, Huey J, Anderson B, Magera MJ, Matern D, Rinaldo P, Robinson BH, Cameron JM, Hahn SH (2005) Automated spectrophotometric analysis of mitochondrial respiratory chain complex enzyme activities in cultured skin fibroblasts. Clin Chem 51(11):2110–2116 10. Karkucin´ska-Wie˛ckowska A, Czajka K, Wasilewski M, Sykut-Cegielska J, Pronicki M,
Pronicka E, Zabłocki K, Duszyn´ski J, Wie˛ckowski MR (2006) Blue native electrophoresis: an additional useful tool to study deficiencies of mitochondrial respiratory chain complexes. Ann Diagn Pediatr Pathol 10(3–4):89–92 11. Lebiedzinska M, Duszynski J, Wieckowski MR (2008) Application of “blue native” electrophoresis in the studies of mitochondrial respiratory chain complexes in physiology and pathology. Postepy Biochem 54:217–223 12. Piekutowska-Abramczuk D, Magner M, Popowska E, Pronicki M, Karczmarewicz E, Sykut-Cegielska J, Kmiec T, Jurkiewicz E, Szymanska-Debinska T, Bielecka L, Krajewska-Walasek M, Vesela K, Zeman J, Pronicka E (2009) SURF1 missense mutations promote a mild Leigh phenotype. Clin Genet 76(2):195–204 13. Pronicki M, Sykut-Cegielska J, Matyja E, Musialowicz J, Karczmarewicz E, Tonska K, Piechota J, Piekutowska-Abramczuk D, Kowalski P, Bartnik E (2007) G8363A mitochondrial DNA mutation is not a rare cause of Leigh syndrome - clinical, biochemical and pathological study of an affected child. Folia Neuropathol 45(4):187–191 14. Pronicki M, Matyja E, PiekutowskaAbramczuk D, Szymanska-Debinska T, Karkucinska-Wieckowska A, Karczmarewicz E, Grajkowska W, Kmiec T, Popowska E, Sykut-Cegielska J (2008) Light and electron microscopy characteristics of the muscle of patients with SURF1 gene mutations associated with Leigh disease. J Clin Pathol 61 (4):460–466 15. Seligman AM, Karnovsky MJ, Wasserkrug HJ, Honker JS (1968) Non-droplet ultrastructural demonstration of cytochrome oxidase activity whit polymerizing osmiophilic reagent, DAB. J Cell Biol 38:1
Chapter 7 An Update on Isolation of Functional Mitochondria from Cells for Bioenergetics Studies Magdalena Lebiedzinska-Arciszewska, Lech Wojtczak, and Mariusz R. Wieckowski Abstract Mitochondria are the organelles where the most fundamental processes of energy transformation within the cell are located. They are also involved in several processes like apoptosis and autophagy, reactive oxygen species formation, and calcium signaling, which are crucial for proper cell functioning. In addition, mitochondrial genome hosts genes encoding important proteins incorporated in respiratory chain complexes and indispensable for the oxidative phosphorylation. Studying isolated mitochondria is, therefore, crucial for better understanding of cell physiology. The presented protocol describes a relatively simple and handy method for crude mitochondrial fraction isolation from different mammalian cell lines. It includes mechanical cells disruption (homogenization) and differential centrifugation. In addition, this chapter presents two basic ways to assess mitochondrial functionality: by measuring mitochondrial inner membrane potential and coupled respiration. Key words Mitochondria isolation, Cell cultures, Oxygen consumption, Mitochondrial membrane potential, Oxidative phosphorylation
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Introduction Methods of mitochondrial fraction isolation from organs like liver, heart, brain, and skeletal muscles have been already published in late 40s of the last century (e.g., (1)). Later on, they have been improved and adapted to different materials and experimental conditions. They are based on the mechanical disintegration of the tissue by homogenization or grinding with rotating blades (Polytron), sometimes with the help of proteolytic enzymes (e.g., trypsin or nagarse), followed by the differential centrifugation (2–5). Isolation of mitochondria from cells naturally living in suspensions, as
Lech Wojtczak deceased 30th September, 2019 Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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lymphocytes, or from adherent cell cultures, is complicated by the fact that some types of cells are more resistant to mechanic disruption even than cells of solid tissues like skeletal muscles. To overcome this problem, homogenization step is often being preceded, or even replaced, by the plasma membrane disruption with the use of heparin (6), digitonin (7), or nitrogen cavitation (8). Alternatively, depending on the experimental requirements, cells can be sonicated (9). Nevertheless, the classical homogenization remains the most universal procedure. It is mostly performed using PotterElvehjem type homogenizers either manually driven with a glass or Teflon pestle or a motor-driven with a Teflon pestle. Though manually driven, a glass pestle seems to be more efficient than a Teflon one in the rupture of cell culture-derived material in our practice. Regardless of cells disruption technique, mitochondrial fraction needs to be separated from the cytosol and other organelles (nuclei, endoplasmic reticulum (ER), etc.). The most common and simplest method to obtain mitochondrial fraction is differential centrifugation. The result of the basic homogenate centrifugation is a pellet of crude mitochondria and a supernatant containing other organelles. Further purification can be achieved by the density-gradient centrifugation. The protocol for highly purified mitochondrial fraction isolation, suitable for proteomic studies but not proper for studying bioenergetics parameters, has been refined and described by our group as well (10). This method involves Percoll gradient ultracentrifugation, which separates mitochondria from interacting fraction of ER, namely mitochondria-associated membranes, or from the other organelles remains, which are detectable in the crude mitochondrial fraction. An alternative procedure to separate mitochondria proposed by Hornig-Do et al. (11) uses magnetic microbeads coated with an antibody against one of the mitochondrial outer membrane proteins (TOM22). Nowadays, the immunoprecipitation-based protocol of mitochondria isolation is more popular, and proper materials are commercially available (11, 12). However, still the centrifugation-based method is faster and more suitable for functional studies. Some latest protocols are also based on the commercially available mitochondria isolation kits (13–15). If the mitochondrial fraction is going to be checked for the functional parameters, it is extremely important to choose the right kit, which is dedicated to this purpose. Particular attention should be payed to the detergents, potentially present in buffers supplied with the kit, or whether the conditions of the preparation aren’t too stringent, what could affect mitochondrial integrity. It should be mentioned that for certain purposes, it is not necessary for mitochondria to be extracted out of the cells as a separated fraction. As an example, our group has shown that the
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certain properties of mitochondria like coupled respiration and membrane potential formation can be studied within the cells after their controlled permeabilization with digitonin (so-called “mitochondria in situ”) (16). A comprehensive description of modern methods for mitochondria isolation from a broad variety of biological material, including cultured cells, has been published by Palotti and Lenaz (17) in 2007 and revised later by other authors (18, 19). The present methodical chapter describes a simple procedure to isolate crude fraction of functional mitochondria from various cell cultures (growing either in suspension or adherent) using homogenization and differential centrifugation.
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Materials
2.1 Isolation of Crude Mitochondrial Fraction from Cell Lines 2.1.1 Solutions and Chemicals
1. Homogenization buffer: 75 mM sucrose (see Note 1), 225 mM mannitol, 0.1 mM EGTA, and 30 mM Tris–HCl, pH 7.4. Check the pH of the solution when it is already cooled down with pH meter and adjust if necessary (see Note 2). Store at 4 C (see Note 3). 2. Mitochondria isolation buffer: 75 mM sucrose (see Note 1), 225 mM mannitol, and 5 mM Tris–HCl, pH 7.4. Check the pH of the solution when it is already cooled down with pH meter and adjust if necessary (see Note 2). Store at 4 C (see Note 3). 3. Proteases and phosphatases inhibitors may be added in prior to proteomic analysis. Avoid supplementation with inhibitors at this step when functional studies are going to be performed.
2.1.2 Equipment and Accessories
1. Motor stirrer with speed controller. 2. pH meter. 3. Motor-driven Potter-Elvehjem homogenizer with tightly fitting Teflon pestle. 4. Potter-Elvehjem homogenizer with a loose fitting glass or Teflon pestle. 5. Centrifuge with temperature control and proper tubes.
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Cell Culture
2.2.1 Materials and Accessories
1. Cell line of interest, for example, HeLa, Mouse Embryonal Fibroblasts (MEF) or Neonatal Human Dermal Fibroblasts (NHDF), Jurkat (T cell leukemia cell line), or rat hepatoma (AS-30D). 2. Cell culture dishes or flasks for lines growing in monolayer and in suspension, respectively. 3. Cell scrapers, tubes. 4. Centrifuge.
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2.2.2 Media and Solutions
1. Choosing the proper culture medium follows the instructions provided by the cell line supplier. Typically, fibroblasts grow in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum, 2 mM L-glutamine, and 1.2% penicillin-streptomycin solution. 2. Phosphate-buffered saline (PBS), without Ca2+ and Mg2+. 3. Trypsin-EDTA solution.
2.3 Determination of Protein Concentration 2.3.1 Equipment, Accessories and Reagents
2.4 Measurement of the Mitochondrial Transmembrane Potential 2.4.1 Solutions and Reagents
1. Spectrophotometer UV/Vis or a plate reader (e.g., Tecan Infinite). Proper acryl cuvettes (1.5 ml) or 96-well transparent plate. 2. Protein Assay Dye Reagent Concentrate (Bio-Rad). 3. Bovine serum albumin standard solution, stock concentrated 1 mg/ml in deionized water. 1. 100 mM phosphate buffer pH 7.4: directly before use mix 4.05 ml of Na2HPO4 solution (1.42 g of Na2HPO4 dissolved in 50 ml of deionized water) with 0.95 ml of NaH2PO4 solution (1.39 g NaH2PO4 dissolved in 50 ml of deionized water) and add 5 ml of deionized water to the final volume of 10 ml. 2. Measurement medium: 75 mM sucrose, 225 mM mannitol, 1 mM MgCl2, 1 mM phosphate buffer (prepared as described above), 0.5 mM EGTA, and 25 mM Tris–HCl, pH 7.4. Control and adjust the pH of the cooled down buffer. Store at 4 C for up to 2 weeks. 3. Safranin O 5 mM aqueous stock solution. 4. Glutamate stock: 0.5 M glutamate aqueous solution, pH 7.4 (adjusted with KOH). 5. Malate stock: 0.5 M malate aqueous solution, pH 7.4 (adjusted with KOH). 6. Rotenone stock: 1 mM rotenone ethanol solution. 7. Succinate stock: 0.5 M succinate aqueous solution, pH 7.4 (adjusted with KOH). 8. CCCP stock: 1 mM CCCP ethanol solution.
2.4.2 Equipment and Accessories
1. Spectrofluorometer or a plate reader (e.g., Tecan Infinite). 2. Acryl cuvettes 10 10 48 mm for spectrofluorometer or a transparent 24-well plate suitable for plate reader.
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2.5 Measurement of the Oxygen Consumption 2.5.1 Solutions and Reagents
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1. 100 mM phosphate buffer pH 7.4: directly before use mix 4.05 ml of Na2HPO4 solution (1.42 g of Na2HPO4 dissolved in 50 ml of deionized water) with 0.95 ml of NaH2PO4 solution (1.39 g NaH2PO4 dissolved in 50 ml of deionized water) and add 5 ml of deionized water to the final volume of 10 ml. 2. Measurement medium: 75 mM sucrose, 225 mM mannitol, 1 mM MgCl2, 1 mM phosphate buffer, 25 mM Tris–HCl, pH 7.4. Control and adjust the pH of the cooled down buffer. Store at 4 C for up to 2 weeks. 3. Glutamate stock: 0.5 M glutamate aqueous solution, pH 7.4 (adjusted with KOH). 4. Malate stock: 0.5 M malate aqueous solution, pH 7.4 (adjusted with KOH). 5. ADP stock: 100 mM ADP aqueous solution. 6. Oligomycin stock: 1 mM oligomycin ethanol solution.
2.5.2 Equipment
1. Clark-type oxygen electrode (e.g., YSI, Yellow Springs, OH, USA or Oroboros Oxygraph-2 k, Oxygraph, Bioenergetics and Biomedical Instruments, Innsbruck, Austria) equipped with an oxygen consumption rate-calculating unit (first derivative of the oxygen concentration trace). 2. Separated injectors for the selected compounds.
3
Methods To isolate an amount of mitochondria sufficient to perform all designed functional studies, the number of plates or flasks needed to be prepared should be experimentally defined based on the cell culture properties. Typically to isolate sufficient amount of mitochondria from fibroblasts growing in monolayer, 30–60 of confluent 10 cm dishes are necessary. Less material is usually needed to isolate crude mitochondrial fraction for basic proteomic studies like western blot. Measurements of the mitochondrial membrane potential (ΔΨ) and oxygen consumption are often used to verify the quality of mitochondrial preparations. The purity of mitochondria and the integrity of the outer mitochondrial membrane can be verified by immunochemical (western blot) analysis by the presence of typical proteins for each of the samples: mitochondrial fraction and for the sample of supernatant from the centrifugation dividing the mitochondria from cytosolic fraction (Subheading 3.1, step 7). Mitochondrial fraction should be enriched in, for example, cytochrome c, superoxide dismutase 2 (SOD2), Tom20, or HSP60 and deprived of, for example, GAPDH. The opposite situation should be observed in cytosol containing fraction.
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3.1 Isolation of Crude Mitochondrial Fraction from Cell Lines
1. Cells Preparation: (a) Adherent Cell Culture-Like MEF: Remove the medium, wash the cells twice with PBS (without Ca2+ and Mg2+), and detach cells by the treatment with trypsin solution for 3–5 min in the incubator (37 C). Add culture medium (e.g., DMEM) to stop trypsinization and centrifuge cells containing suspension for 3 min at 200 g at room temperature. Afterwards, wash the pellet twice by gentle resuspending in PBS and centrifuging. (b) Alternatively, the adherent cell culture may be removed with the use of cell scraper after a brief wash with PBS, to avoid trypsinization, which may additionally stress the cells and affect mitochondrial parameters. Then, the cells should be collected in the tube and centrifuged to obtain the cell pellet, which after a wash with homogenization buffer is ready to be processed as described in Subheading 3.1, step 2. (c) Cells Growing in Suspension: Transfer the cell suspension to the tube and centrifuge for 3 min at 200 g at room temperature. Discard the supernatant and resuspend the cells in PBS. Centrifuge for 3 min at 200 g at room temperature. Repeat the washing once again. 2. Resuspend the cell pellet in approximately 20 ml of the ice-cold homogenization medium (see Note 4). All the steps of the mitochondria isolation protocol from now on require low temperature conditions. Therefore, we recommend keeping all the buffers and materials on ice (see Note 5). 3. Homogenize the cells in a tightly fitting Potter-Elvehjem homogenizer with a glass or Teflon pestle. A motor-driven homogenizer can be used to facilitate the homogenization; however, some type of cell pellets are easier to disintegrate by manual homogenization with the use of tight-fitting glass pestle in a glass Potter-Elvehjem homogenizer (see Note 6). 4. Check the integrity of homogenized cells few times during the homogenization procedure with a light microscope by pouring a small drop of the homogenate on a glass microscope slide. The homogenate is ready for further processing when at least 70% of the cells sample is smashed and homogenous.
5. Centrifuge the homogenate for 5 min at 1000 g at 4 C. 6. Discard the pellet (unbroken cells and nuclei) and centrifuge the supernatant again for 5 min at 1000 g at 4 C. 7. Discard the pellet (remaining unbroken cells and nuclei) and centrifuge the supernatant for 10 min at 8000 g at 4 C.
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8. Discard the supernatant (containing: cytosolic fraction, plasma membrane, lysosomes, and microsomes). 9. Gently resuspend the mitochondrial pellet in approximately 10 ml of the mitochondria isolation medium using PotterElvehjem homogenizer with a loose-fitting pestle and centri fuge it again for 10 min at 8000 g at 4 C (see Note 5). 10. Discard the supernatant and gently resuspend the mitochondrial pellet in 5–10 ml (depending on the pellet volume) of the mitochondria isolation medium using Potter-Elvehjem homogenizer with a loose-fitting pestle. Centrifuge it again for 10 min at 10,000 g at 4 C. 11. Gently resuspend the crude mitochondrial pellet in ~2 ml of the mitochondria isolation medium using a loose PotterElvehjem homogenizer and store the sample on ice (see Note 5). The material can now be used for functional studies like measurement of oxygen consumption, mitochondrial membrane potential, production of reactive oxygen species as well as calcium uptake. If the proteomic analysis is scheduled, it might be necessary to perform further purification of this crude mitochondrial fraction in order to obtain a pure mitochondria deprived of the other organelles remains as described in (5). 3.2 Measurement of Protein Concentration
Protein measurement with the use of both Tecan multiwell plate reader and spectrophotometer is based on the Bradford’s method (20). 1. Fill the 1.5 ml spectrophotometer cuvette with 1.2 ml of ddH2O. Add 1–2 μl of the mitochondrial sample to the cuvette (Prepare a proper mitochondrial sample dilution as the final absorbance of the sample should not exceed 0.6). Add 300 μl of Bio-Rad Protein Assay Dye Reagent and shake the sample. Measure the absorbance at 595 nm. Calculate the result based on the BSA standard solution curve including absorbance measurement for at least five points, for example: 1.0, 2.0, 5.0, 10, and 12 μg of BSA protein per cuvette. 2. Prepare the, at least, 10-diluted mitochondrial sample and the dilution of BSA standard. Mark the wells for standard curve for at least five points of BSA concentrations in two repetitions each. Mark the wells for measurement of the protein concentration in the mitochondrial sample. Add 120 μl of the 5-diluted Bio-Rad Protein Assay Dye Reagent in ddH2O to each well and the appropriate BSA or sample dilution, which
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should be determined experimentally to be able to measure absorbance in the appropriate range. Mix gently and read the absorbance at 595 nm then calculate the protein concentration based on the standard curve from the same plate. 3.3 Measurement of the Mitochondrial Membrane Potential with the Use of Safranin O
1. Adjust the spectrofluorometer: excitation wavelength, 495 nm; emission wavelength, 586 nm; slits (excitation and emission), ~3 nm. 2. Fill the fluorometer cuvette with 3 ml of the measurement medium (see Note 7). 3. Add 3 μl of 5 mM safranin O, 30 μl of 0.5 M glutamate, and 30 μl of 0.5 M malate stock solutions. 4. Start measuring fluorescence. 5. Add the mitochondrial suspension corresponding to about 1 mg protein and observe changes in the fluorescence. When it reaches a plateau, continue with the following consecutive additions: 5 μl of 1 mM rotenone, next 30 μl of 0.5 M succinate, and at the end, 5 μl of 1 mM CCCP (to uncouple mitochondria) stock solutions. An example of the results is shown in Fig. 1a. Alternatively, the mitochondrial membrane potential can be measured with the use of multiwell plate reader like Tecan Infinite. It requires adjustable wavelength or proper filters for excitation, 495 nm and emission, 586 nm. The sensitivity of the scan should be experimentally defined as well as the parameters of the kinetic measurement. 1. Adjust excitation wavelength, 495 nm; emission wavelength, 586 nm and gain (optimal gain should be defined experimentally to operate in a nonsaturated fluorescence signal range). 2. Fill the 24-multiplate wells with 1 ml of the measurement medium (75 mM sucrose, 225 mM mannitol, 1 mM MgCl2, 1 mM phosphate buffer, 0.5 mM EGTA, and 25 mM Tris– HCl, pH 7.4) containing additionally 5 mM safranin O, 5 mM glutamate, and 5 mM malate (see Note 7). 3. Start measuring fluorescence. 4. Add the mitochondrial suspension corresponding to about 300 μg protein and observe changes in the fluorescence. When it reaches a plateau, continue with the following consecutive additions: 1 μl of 1 mM rotenone, next 10 μl of 0.5 M succinate, 1 μl of 1 mg/ml antimycin A, next 25 μM TMPD added together with 2 mM ascorbate, and at the end, 5 μl of 1 M azide (to inhibit complex IV) stock solutions. An example of the results is shown in Fig. 1b.
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B CCCP
0
10
2000
ROT
14000
no 16000
Succinate
960
900
840
900
780
600
time [s]
720
300
660
0
TMPD/Ascorbate 600
20000
Mitochondria
540
18000
Succinate
80
12000
480
no
Mitochondria
10000
420
50
max. Azide
8000
360
40
6000
300
30
Antimycin A
4000
240
20
180
Fluorescence Intensity
Fluorescence intensity
max.
120
ROT
0
4
60
A
87
time [s]
Fig. 1 Measurement of mitochondrial membrane potential (ΔΨ) with the use of safranin O in mitochondria isolated from HeLa cells (a) ΔΨ measured with the use of spectrofluorometer. Addition of rotenone (ROT, in the presence of glutamate and malate as substrates) leads to the collapse of ΔΨ due to the inhibition of complex I of the mitochondrial respiratory chain. Subsequent addition of succinate (complex II substrate) restores ΔΨ. Addition of CCCP (a protonophore) leads to the collapse of ΔΨ (uncoupling effect) (b) ΔΨ measured with the use of multiwell plate reader. Addition of rotenone (ROT, in the presence of glutamate and malate as substrates) leads to the collapse of ΔΨ due to the inhibition of complex I of the mitochondrial respiratory chain. Subsequent addition of succinate (complex II substrate) restores ΔΨ. Addition of antimycin A (complex III inhibitor) leads to the collapse of ΔΨ. Subsequent addition of TMPD/ascorbate (complex IV substrate) restores ΔΨ. Addition of azide (complex IV inhibitor) leads to the collapse of ΔΨ again 3.4 Measurement of Oxygen Consumption (Respiration)
1. Fill the chamber with the measurement medium (see Note 7). 2. Add 10 μl of 0.5 M glutamate and 10 μl of 0.5 M malate stock solutions. 3. Start running the oxygen concentration trace. 4. Add mitochondrial suspension (approximately 1 mg protein). 5. Wait for a stable signal (usually it takes about 1–3 min). Then, continue the measurement with the following consecutive additions: 5 μl of 100 mM ADP and later 2 μl of 1 mM oligomycin stock solutions. An example of the results is shown in Fig. 2.
4
Notes 1. To isolate intact mitochondria, it is necessary to use low-calcium sucrose (e.g., Merck, cat. no. 100892.9050). 2. To adjust the pH, use KOH (avoid NaOH) or HCl. 3. Homogenization medium prepared in advance should be kept at 4 C for maximum 2 weeks.
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Mitochondria O2 concentration (nmol/ml)
ADP 240 220 200
O2 concentration 180
Oligo rate of oxygen consumption (first derivative of the O2 concentration trace)
160
Time Fig. 2 Evaluation of the oxygen consumption rate in mitochondria isolated from HeLa cells Mitochondrial respiration at low ADP level (state 4) is slow but increases dramatically after addition of exogenous ADP (state 3). Addition of oligomycin (oligo, inhibitor of ATP synthase) results in a decrease in oxygen consumption due to the inhibition of ATP production. Respiratory control ratio (RCR) is the quotient of state 3 and state 4. High RCR value indicates better integrity of the inner mitochondrial membrane and tighter coupling between the electron transport and ATP synthesis
4. Homogenization and the following steps must be performed at 4 C to minimize the activity of proteases, phosphatases, and phospholipases. All solutions should be refrigerated at 4 C and all equipment precooled. 5. Extreme care should be taken to avoid contamination with the ice and tap water. 6. Pay attention to the homogenizer heating during the homogenization procedure. Do not exceed with the speed of the motor-driven homogenizer and the time of the procedure. 7. Prior to perform the assays, the measurement media should be at the room temperature.
Acknowledgments This work was supported by National Science Center grant no. 2015/17/D/NZ1/00030 for ML-A and by the Polish National Science Centre grant (UMO- 2018/29/B/NZ1/ 00589) for MRW.
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References 1. Hogeboom GH, Schneider WC, Pallade GE (1948) Cytochemical studies of mammalian tissues. I. Isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate material. J Biol Chem 172:619–635 2. Garcia-Cazarin ML, Snider NN, Andrade FH (2011) Mitochondrial isolation from skeletal muscle. J Vis Exp (49). http://www.jove. com/details.php?id¼2452, https://doi.org/ 10.3791/2452 3. Storrie B, Madden EA (1990) Isolation of subcellular organelles. Methods Enzymol 182:203–225 4. Djafarzadeh S, Jakob SM (2017) Isolation of intact mitochondria from skeletal muscle by differential centrifugation for high-resolution respirometry measurements. J Vis Exp (121): e55251. https://doi.org/10.3791/55251 5. Kras KA, Willis WT, Barker N, Czyzyk T, Langlais PR, Katsanos CS (2016) Subsarcolemmal mitochondria isolated with the proteolytic enzyme nagarse exhibit greater protein specific activities and functional coupling. Biochem Biophys Rep 6:101–107 6. Nessi P, Billesbolle S, Fornerod M, Maillard M, Frei J (1977) Leucocyte energy metabolism. VII. Respiratory chain enzymes, oxygen consumption and oxidative phosphorylation of mitochondria isolated from leucocytes. Enzyme 22:183–195 7. Moreadith RW, Fiskum G (1984) Isolation of mitochondria from ascites tumor cells permeabilized with digitonin. Anal Biochem 137:360–367 8. Kristia´n T, Hopkins IB, McKenna MC, Fiskum G (2006) Isolation of mitochondria with high respiratory control from primary cultures of neurons and astrocytes using nitrogen cavitation. J Neurosci Methods 152(1–2):136–143 9. Gellerfors P, Nelson BD (1979) A rapid method for the isolation of intact mitochondria from isolated rat liver cells. Anal Biochem 93:200–203 10. Wieckowski MR, Giorgi C, Lebiedzinska M, Duszynski J, Pinton P (2009) Isolation of mitochondria-associated membranes and mitochondria from animal tissues and cells. Nat Protoc 4:1582–1590 11. Hornig-Do HT, Gu¨nther G, Bust M, Lehnartz P, Bosio A, Wiesner RJ (2009)
Isolation of functional pure mitochondria by superparamagnetic microbeads. Anal Biochem 389:1–5 12. Franko A, Baris OR, Bergschneider E, von Toerne C, Hauck SM, Aichler M, Walch AK, Wurst W, Wiesner RJ, Johnston IC, de Angelis MH (2013) Efficient isolation of pure and functional mitochondria from mouse tissues using automated tissue disruption and enrichment with anti-TOM22 magnetic beads. PLoS One 8(12):e82392 13. Pashkovskaia N, Gey U, Ro¨del G (2018) Mitochondrial ROS direct the differentiation of murine pluripotent P19 cells. Stem Cell Res 30:180–191 14. Gao J, Liu S, Xu F, Liu Y, Lv C, Deng Y, Shi J, Trilobatin GQ (2018) Protects against oxidative injury in neuronal PC12 cells through regulating mitochondrial ROS homeostasis mediated by AMPK/Nrf2/Sirt3 signaling pathway. Front Mol Neurosci 11:267 15. Rambold AS, Kostelecky B, Elia N, LippincottSchwartz J (2011) Tubular network formation protects mitochondria from autophagosomal degradation during nutrient starvation. Proc Natl Acad Sci U S A 108:10190–10195 16. Bogucka K, Wroniszewska A, Bednarek M, Duszyn´ski J, Wojtczak L (1990) Energetics of Ehrlich ascites mitochondria: membrane potential of isolated mitochondria and mitochondria within digitonin-permeabilized cells. Biochim Biophys Acta 1015:503–509 17. Pallotti F, Lenaz G (2007) Isolation and subfractionation of mitochondria from animal cells and tissue culture lines. Methods Cell Biol 80:3–44 18. Frezza C, Cipolat S, Scorrano L (2007) Organelle isolation: functional mitochondria from mouse liver, muscle and cultured fibroblasts. Nat Protoc 2(2):287–295 19. Wettmarshausen J, Perocchi F (2017) Isolation of functional mitochondria from cultured cells and mouse tissues. Methods Mol Biol 1567:15–32 20. 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
Chapter 8 PCR-Based Determination of Mitochondrial DNA Copy Number in Multiple Species Tess C. Leuthner, Jessica H. Hartman, Ian T. Ryde, and Joel N. Meyer Abstract Mitochondrial DNA (mtDNA) copy number is a critical component of overall mitochondrial health. In this chapter, we describe methods for simultaneous isolation of mtDNA and nuclear DNA (nucDNA), and measurement of their respective copy numbers using quantitative PCR. Methods differ depending on the species and cell type of the starting material, and availability of specific PCR reagents. We also briefly describe factors that affect mtDNA copy number and discuss caveats to its use as a biomarker. Key words Mitochondrial DNA, mtDNA, mtDNA depletion, Copy number, QPCR, Mitochondrial toxicity, Mitochondrial disease
1
Introduction In humans and most animals, the mitochondrial genome (mtDNA) encodes 13 proteins, all of which are components of the electron transport chain (ETC) or ATP synthase complexes and are essential for oxidative phosphorylation (OXPHOS) [1]. On average, each cell contains between 103 and 104 copies of the mitochondrial genome, though this number varies by orders of magnitude between cell type and developmental stage. The processes that maintain this variation within and between individuals are still under investigation [2], making mtDNA copy number (mtDNA CN) a fascinating area of research across many fields of study. mtDNA CN is regulated by coordination of mtDNA replication and removal [3, 4]. mtDNA replication is carried out independently of the cell cycle by the nuclear DNA (nucDNA)-encoded polymerase γ, the only replicative DNA polymerase thought to be normally found in the mitochondria [5, 6]. mtDNA replication and mitochondrial biogenesis (to which it appears to be closely linked under most circumstances) are additionally regulated in part by
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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mitochondrial transcription factor A (TFAM) [7] and other molecular factors [8, 9]. mtDNA CN is also regulated by the removal of mtDNA via mitochondrial degradation by a number of mechanisms. The best understood of these are autophagy and mitophagy [10]. In addition, at least under some circumstances, cellular import and export of mitochondria (likely including mtDNA, although this has not been demonstrated in all cases) to neighboring cells [11–13] or into circulation may occur [14, 15]. Presumably as a result of these processes, mtDNA CN can be measured in extracellular samples including human serum [16, 17] and urine, where it has mostly been used as a marker of kidney injury [18, 19]. It should be noted that the source of circulating and urinary mtDNA is not easy to determine. Maintenance of mtDNA CN is critical to health and disease, as illustrated by the fact that dysregulation of copy number can manifest in many diseases. For example, mutations in nine genes involved in mtDNA replication and nucleotide metabolism cause mitochondrial DNA depletion syndrome (MDS) in humans [20]. There are multiple, rare diseases associated with MDS that often present in young children, such as Alper’s syndrome, as well as other recessive neuropathies and myopathies [20]. mtDNA depletion is also implicated in more common diseases including type 2 diabetes [21], many cancers [22], and neurodegenerative disorders such as Parkinson Disease [5, 23], though direct causal links have not yet been established. Pharmaceuticals can also block mtDNA replication and result in mtDNA depletion, as in the case of the nucleoside reverse transcriptase inhibitors (NRTIs), which are used to treat human immunodeficiency virus (HIV). NRTIs are nucleoside analogs that inhibit mtDNA replication and can cause toxicity mediated in part by mtDNA depletion [24]. Furthermore, growing evidence suggests that environmental exposures can alter mtDNA CN [4, 25, 26]. Interestingly, the direction and magnitude in which mtDNA CN changes (especially in epidemiological studies) remain somewhat contradictory. For example, one population-based exome-wide association study showed an increase in mtDNA CN associated with cigarette smoking, but high PM2.5 was associated with a decrease in mtDNA CN [27]. Another study demonstrated that leukocyte mtDNA CN was inversely related to PM2.5 exposure [28]. Hou et al. (2015) also found an inverse association of mtDNA CN in blood samples from individuals exposed to increasing levels of traffic exhaust exposure; while on the other hand, Zhong et al. (2016) found increased mtDNA CN in individuals with higher traffic exhaust exposure [29, 30]. Pavanello et al. (2013) also found an increase in mtDNA CN in peripheral blood lymphocytes in individuals with higher exposures to polycyclic aromatic hydrocarbons [31].
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Laboratory models have also been used to understand the effect of the environment on mtDNA CN. For example, a recent study in rats reported an increase in mtDNA CN in liver tissue after exposure to perfluorooctanoic acid [32]. Cell culture studies have similarly shown that exposures to a number of chemicals can either increase or decrease mtDNA CN [33–36]. In C. elegans, we found that an early-life exposure to a low dose of ultraviolet C radiation results in decreased mtDNA CN [37], but we have not yet observed a change in copy number with other chemical exposures in this model [38, 39], with the exception of the well-known mtDNA replication inhibitor ethidium bromide [40]. Finally, there is strong evidence that nonchemical environmental factors including exercise [41–43], diet [44–46], age [47–49], and obesity [50, 51] also modulate mtDNA CN. More recent evidence also suggests an association between psychosocial stress and altered mtDNA CN [52]. Interpretation of mtDNA CN is complicated by the fact that mtDNA CN results from a balance of biogenesis and removal. Thus, alterations compared to a control or reference group may result from a perturbation in either biogenesis or removal, or both. As a result, stressors may result in either increases or decreases in mtDNA CN under many different exposure scenarios, as previously discussed theoretically [4] and demonstrated empirically in the case of doxorubicin [4, 35]. Perhaps for this reason, as illustrated by the examples provided above, there is evidence for both increases (stress-induced, presumably as an attempt at adaptation, or inhibition of turnover, such as mitophagy) and decreases (presumably reflecting either removal of damaged mitochondria or inhibition of mtDNA replication) associated with exposures in both the epidemiological and experimental literature. Furthermore, as mentioned, mtDNA CN is affected by a wide range of environmental and genetic factors. This means that changes in mtDNA CN associated with environmental and other factors should be interpreted with care, and mtDNA CN is unlikely to be a specific biomarker of any given disease or stressor. It is possible, however, that it may serve as one part of a panel of biomarkers and as a way to understand the potential role of mitochondrial biology in specific contexts. For example, we found in a small study that mtDNA CN was elevated in veterans with Gulf War illness and interpreted those results not to mean that this elevation is a specific biomarker of GWI, but rather as support for mitochondrial dysfunction or dysregulation in the context of this disease [53]. This chapter provides protocols for the simultaneous isolation of mitochondrial and nuclear DNA, and measurement of both genome copy numbers from a variety of species. We present two protocols for copy number determination: real-time and non-realtime qPCR. They differ based on availability and optimization of
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reagents. In C. elegans and human samples, absolute mtDNA CN can be measured via real-time qPCR, using a plasmid-based mtDNA CN standard curve. In Danio rerio, Fundulus heteroclitus, Mus musculus, and Rattus norvegicus, real-time qPCR can also be used, however, without a standard curve (until these are developed), such that the measurement of mtDNA CN is expressed as a ratio, relative to nuclear DNA CN. Nuclear DNA CN is a good proxy for cell number except in multinucleate cell types. The primer sequences and details regarding specific PCR parameters for each species are located in Table 1 (real-time qPCR) and Table 2 (non-real-time qPCR) below.
2 2.1
Materials DNA Isolation
2.1.1 C. elegans, Small Number
1. Worm Lysis Buffer: Standard 3.3 PCR buffer, nuclease-free H2O, 20 mg/mL proteinase K. 2. Platinum worm pick. 3. Thin-walled PCR tubes. 4. Thermal cycler or heat block. 5. Dry ice and/or -80 C freezer. 3.3 Worm Lysis Buffer (40 ml)—Adapted from (Cheng, 2001): l 82.5 mM Tricine (pH 8).
2.1.2 C. elegans, Large Numbers or Animal Tissue
l
264 mM potassium acetate.
l
36.2% w/v glycerol.
l
7.425% v/v DMSO.
l
16.95 ml nuclease-free water.
l
Pipette into 1 mL aliquots.
l
Store at 20 C indefinitely.
1. K medium: 31.5 mM KCl, 51 mM NaCl in ddH2O (C. elegans). 2. RNAlater solution (animal tissue). 3. 15 mL conical tubes. 4. Liquid nitrogen. 5. Dry ice. 6. Mortar and pestle (C. elegans and tough animal tissue such as muscle). 7. Handheld homogenizer (softer animal tissue such as liver). 8. Qiagen G/20 Genomic Tips Kit. 9. Isopropanol.
nuc
F. heteroclitus mt
nuc
nd1
R. norvegicus mt
CFTR
16S rRNA
Reverse primer Seq 50 -3’
CGT TTA CCC CAG ATG CAC CT TGG ATA CCT GAC CGA GAG CT
CTA GCA GAA ACA AAC CGG GC GCC AGC CTC TCC TGA TTT TAG TGT
CAC CCA AGA ACA GGG TTT GT TGC TGT CTC CAT GTT TGA TGT ATC T
AAA ATT AAC GGC CCC AAC CC GCC GCT GCC TTC ATT GCT GT
86
107
164
75
CCG AGT TCC TTC TTC CCC TT ATG AGC TGG GTG TGC GCT GA
AGG CGT TCT GAT GAT GGG AA GTT TGC TTG CCG ACT CCT TG
GTG CGA TTG GTA GGG CGA TA AGA CAA CTC TTA CGG CTG GC
234
131
144
181
90
195
60
60
62
60
60
60
60
60
62
62
60
60
Gonzalez-Hunt et al. (2016) [58]
Gonzalez-Hunt et al. (2016) [58]
Gonzalez-Hunt et al. (2016) [58]
Quiros et al. (2017) [57]
Venegas and Halberg (2012) [56]
Rooney et al. (2015) [55]
Bratic et al. (2009) [54]
Amplicon Annealing (bps) temperature Reference
CCG GCT GCG TAT – TCT ACG TT GGG AAC ACA AAA – GAC CTC TTC TGG
TGG CCA TGG GTA TGT TGT TA TCT CTG CTC CCC ACC TCT AAG T
AGC GTC ATT TAT AAG CTT GTG CTA TGG GAA GAA GAC ATC CCA TAA ATG T GCC GAC TGG AAG GCG GAG ATC ACC AAC TTG TC TTC CAG TA
Forward primer Seq 50 -3’
CAA ACC TTT CCT GCA CCT CC 30 ,50 -cyclic AMP GTT CCC GCC TTC TTC CTC TG phosphodiesterase (PDE41, PDE4-2)
vtg2
nuc
hk2
nuc
nd1
nd1
mt
B2-microglobulin
nuc
mt
D. rerio
M. musculus
tRNA-Leu(UUR)
cox-4
nuc
mt
nd-1
mt
C. elegans
H. sapiens
Genome Target gene
Species
Table 1 Real-Time PCR primers and conditions
mtDNA Copy Number Determination 95
mt
mt
M. musculus
R. norvegicus
H. sapiens
C. elegans
D. rerio
Reverse primer Seq 50 -30
Previously published nd-6 primersc
nd-5
pole-1
nuc
ahr-2
nuc
mt
16S rRNA
233
198
63
63
60
62
65
152
195 TGT CCT CAA GGC TAC CAC CTT CTT CA TCC CGT CTA TTG GAC GCG CAC GAT 225 CAG GTC TTT CCA ATC TCG ATT TTC CAC ACC GGT GAG GTC TTT GGT TC
CAA ACA CAA GCC CAC TGA CTT GAT TCG CCT GTT TAC GGG GGA GAC AGT ATG GGC TGG GCG ACA TGT GCA TGT ATA AAA TTG G CGC TCC CAA A
61
151
60
23
18
27
21
24
19
21
Hunter et al. (2010) [59], Meyer et al. (2007) [62] Meyer et al. (2007) [62], Boyd et al. (2010) [63]
Hunter et al. (2010) [59]
Hunter et al. (2010) [59]
Ayala-Torres et al. (2000) [61]
Ayala-Torres et al. (2000) [61]
Ayala-Torres et al. (2000) [61]
Amplicon Annealinga Cyclea (bps) temperature number Reference
211 GTC TGG GTC TCC CCT CCC ATT CAT TAG TAG GTC TGG TAT CGC CGC CCT GAA TGC
Previously published nd-6 primersb
Forward primer Seq 50 -30
GCT CCT GAT ATA CAT GAG CAA TTC GCA TTC CCA CGA CAG CGG ATA AA GTC ACT TCT TGT Ribosomal CGA GGG ATA CCT GTG AGC AGC TT GCT GCC ATC GT protein L11
Cox-1
nd-6
cyt b
mt
nuc
D. melanogaster mt
mt
Species
nd-6
Target Genome gene
Table 2 Quantitative, non-Real-Time PCR primers and conditions
96 Tess C. Leuthner et al.
ATPase subunit 9
CFTR
nuc
mt
12S rRNA
tRNALeu – nd-5 VDAC2
mt
nuc
mt
TGG AGC AGG TAT CTC AAC AA
TTT ACA CAT GCA AGT ATC CG GCC GCT GCC TTC ATT GCT GT
CTC ACA AAC ATC TTT GCA CTC AG
AAC TCC AAG TAG CAG CTA TGC AC
TGT AGC TTC TGA TAA GGC GA
CCG AAG GCT ATC AAC TTG AG ATG AGC TGG GTG TGC GCT GA
AGA ACC TCT CTC CAA AAC ATT CC
GAG GGG TAG AAG GCT TAC AAA AA
158
234
247
140
184
60
62
55
57
59
19
25
25
26
22
Karthikeyan et al. (2003) [64]
Rooney et al. (2015) [55]
Rooney et al. (2015) [55]
b
Annealing temperature and cycle number are suggested starting points and may need optimization based on differences in laboratory equipment and PCR kit used Note: These primers blast with a single mismatch to a nuclear target on chromosome 4 of roughly the same size. Although they may not amplify with the same efficiency, we have concerns that there could be a lack of specificity in some conditions c Note: These primers blast with a single mismatch to a nuclear target on chromosome 5 of roughly the same size. Although they may not amplify with the same efficiency, we have concerns that there could be a lack of specificity in some conditions
a
S. cerevisiae
F. grandis
O. latipes
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10. 70% ethanol. 11. Glass Pasteur pipettes. 12. 1.7 mL microcentrifuge tubes. 13. 50 C water bath. 14. Refrigerated microcentrifuge. 15. Tabletop centrifuge with 15 mL conical tube buckets. 2.1.3 Cultured Cells
1. Either the Qiagen G/20 Genomic Tips Kit and associated buffers (see above) or. 2. A QIAcube for automated DNA isolation, with the QIAmp DNA Mini Kit for human samples or the DNeasy Blood and Tissue Kit (Qiagen) for animal samples. 3. Pellets of approximately 1 106 cells.
2.2 DNA Quantification (See Note 1)
1. PicoGreen dsDNA quantification reagent. 2. Lambda/HindIII DNA standard curve. 3. 1 TE Buffer: 10 mM Tris–HCL, pH 8.0, 1 mM EDTA. 4. Fluorescent plate reader with excitation filter at 480 nm and an emission filter at 520 nm (485 nm and 528 nm also work well). 5. Black or white bottom 96-well plate.
2.3
Real-Time PCR
1. Power SYBR Green PCR Master Mix. 2. Standard 96-well PCR plate with optically clear sealing film. 3. Real-Time PCR System (ABI 7300), or adapt and QC with a similar instrument. 4. ABI Prism 7300 Sequence Detection Software or similar. 5. Primers, species, and target genome-specific, see Table 1. 6. Nuclease-free H2O.
2.4 Quantitative, Non-Real-Time, PCR
1. Standard thermal cycler. 2. KAPA Long-Range Hot Start DNA Polymerase Kit (KAPABiosystems) (optimized for human samples, see Note 2), or. 3. GoTaq Flexi PCR Kit (Promega) (optimized for F. grandis samples, see Note 2). 4. 0.2 mL PCR tubes. 5. PCR hood with germicidal lamp for sterilization. 6. Primers, species, and target genome-specific, see Table 2. 7. All materials from Subheading 2.2 DNA Quantification. 8. 0.1 mg/mL bovine serum albumin in nuclease-free H2O. 9. 10 mM dNTPs mix.
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10. Nuclease-free H2O. 11. Dedicated pipettes and sterile aerosol pipette tips for QPCR setup. 12. Different set of pipettes and regular tips for post-PCR analysis. 13. Distinct workstations for setting up and post-PCR analysis (see Note 3).
3 3.1
Methods DNA Isolation
3.1.1 C. elegans, Small Number of Worms
1. Make fresh 1 worm lysis buffer: 65% nuclease-free water, 30% 3.3 lysis buffer, 5% proteinase K (final concentration of 1 mg/mL). 2. Using a platinum worm pick, transfer six individual, L4 stage or later C. elegans into 90μL of 1 worm lysis buffer prealiquoted into thin-walled PCR tubes (see Notes 4 and 5) and freeze on dry ice (or at 80 C) immediately. If using dry ice, once all samples are picked, transfer to 80 C for at least 10 min. This is usually done in triplicate for each sample and data are averaged (see Note 6). 3. Thaw samples, vortex briefly and spin to collect contents at the bottom of the tube. In a standard thermal cycler or heat block, heat to 65 C for 1 h, followed by 95 C for 15 min, and then hold at 8 C. This crude worm lysate will be used as template DNA for the real-time PCR reactions and does not need to be quantified. This lysate can also be used for the non-real-time quantitative PCR, if real-time PCR is not available.
3.1.2 C. elegans, Large Number of Worms or Animal Tissue (Skip Steps 1 and 2)
1. Wash worms off of bacterial plate with K medium into a 15 mL conical tube, pellet at 2200 g for 2 min, remove medium, and resuspend in 10 mL fresh medium. Gently rock tubes for 20 min to allow worms to clear gut contents. Pellet at 2200 g for 2 min and wash 2 with fresh medium. 2. Resuspend worm pellet in a small volume of medium (residual medium left after wash) with a glass pipette and drip worm suspension directly into liquid nitrogen. 3. Frozen worm pellets can be stored at 80 C indefinitely, animal tissue can be stored at -80 C in RNAlater. As a guide, roughly 10–15 mg of F. grandis liver tissue is sufficient for DNA isolation. 4. Grind frozen worm pellets or tough tissue samples to a fine powder in a liquid nitrogen-cooled mortar and pestle (see Note 7). A squeaking sound is heard when worms are sufficiently
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ground. Alternatively, if the tissue is not tough (i.e., liver tissue), it can be manually homogenized in prealiquoted buffer G2 with RNAse A. 5. Scoop the powder into prealiquoted buffer G2 with RNAse A, as per the Qiagen 20/G Genomic Tips Handbook tissue protocol. 6. Follow the Qiagen 20/G Genomic Tips Tissue protocol for DNA isolation. 3.1.3 Cell Culture Samples
1. Standard DNA isolation methods can be used. We routinely use the Qiagen 20/G Genomic Tips Kit, or for automated DNA isolation, the QIAcube with the QIAamp DNA Mini kit or DNeasy Blood and Tissue Kit can be used (see Note 8).
3.2 DNA Quantification
1. DNA from large-scale worm preparations, cultured cells, or animal tissue needs to be quantified prior to real-time or quantitative non-real-time PCR. DNA from the small-scale worm lysis protocol does not. 2. Prepare a DNA concentration standard curve by diluting Lambda/HindIII DNA to 150 ng/μL, 100 ng/μL, 50 ng/ μL, 25 ng/μL, 12.5 ng/μL, and 0 ng/μL in TE buffer (see Note 9). 3. Dilute DNA samples 1:10 in 1 TE buffer (see Note 10). 4. Add 5μL of DNA and 95μL of 1 TE buffer into two duplicate wells of a black or white 96-well plate (suitable for fluorescence measurements). 5. Add 5μL of each Lambda/HindIII standard and 95μL 1 TE into two duplicate wells. 6. The following three steps should be done in low light conditions (see Note 11). Prepare PicoGreen working solution (100μL of working solution needed per well) by adding 5μL PicoGreen reagent per 1 ml TE buffer (see Note 12). 7. Add 100μL PicoGreen working solution to each well and incubate at room temperature in the dark for 10 min. 8. Measure the fluorescence of each sample with excitation at 480 nm and emission at 520 nm. 9. Determine DNA sample concentrations by comparing fluorescence values to those of the standard curve. If the DNA concentrations are far above the range of the curve, redilute the DNA and measure again (see Note 13). 10. Dilute the sample to 3 ng/μL in TE buffer.
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Real-Time PCR
3.3.1 C. elegans Samples with Standard Curve
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1. Prepare the standard curve (see Note 14) as follows: Thaw an aliquot of the mtDNA CN standard curve plasmid (10 million copies/μL) and dilute to 32,000 copies/μL. Serially dilute 1:1 down to 4000 copies/μL. Add 2μL of each dilution and a 0 copies/μL control (TE buffer or water) to separate wells in the 96-well PCR plate to be used in the copy number PCR. A typical standard curve contains 64,000, 32,000, 16,000, 8000, 4000, and 0 copies per well. For nucDNA copy number, use glp-1 lysate instead of a plasmid. Pick 20 individual 24 h post L4 young adults into 40μL of 1 lysis and store at 80 (1567 copies/μL). Lyse immediately before use. Add 40μL of nuclease-free water to lysate; the concentration will now be 784 copies/μL. Serially, dilute this preparation 1:1 until getting a 24.5 copies/μL dilution. We are currently building a nucDNA standard curve plasmid for C. elegans. 2. Proceed with real-time PCR setup as for all other samples.
3.3.2 All Other Real-Time PCR Samples, Without Standard Curve
1. Assemble the PCR reactions as follows: combine 2μL of template DNA (standard curve dilution, worm lysate, or 3 ng/μL isolated DNA), 2μL of mtDNA target-specific primer pair (400 nM final concentration each, see Note 15), 12.5μL Power SYBR Green PCR Master Mix, and 8.5μL H2O in one well of the 96-well PCR plate. Each sample is amplified in triplicate and the data are averaged (see Note 16). Repeat this step in separate wells using nuclear DNA-specific primers. 2. When analyzing a large number of samples with the same primer pair, a master mix is made containing the reagents that are common to all reactions (Power SYBR Green Master Mix, H2O, and primers) and aliquoted into individual reactions (23μL of master mix and 2μL of lysate or template). 3. Cycle in an ABI 7300 Real-Time PCR System (or comparable system) as follows: 50 C for 2 min, 95 C for 10 min, 40 cycles of 95 C for 15 s, and annealing temperature (primer-specific, Table 1) for 60 s. A dissociation curve is also calculated for each sample to ensure presence of a single PCR product.
3.3.3 Quantitative (NonReal-Time) PCR
1. For samples from C. elegans, D. melanogaster, F. grandis, O. latipes, D. rerio, mouse, rat, and human, relative mtDNA content can be measured in a quantitative, nonreal-time, PCR reaction in which the PCR product is quantified after completion of the reaction. Nuclear copy number can also be measured using this protocol, and primers are listed in Table 2 for some nuclear targets, however, this is not often necessary (see Note 17). 2. Extra control reactions are necessary for this protocol. Be sure to include a “50% control” that contains control template DNA
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(or worm lysate) diluted 1:1 with H2O or TE buffer prior to being added to the reaction (it is not advised to simply add ½ the volume of the control sample) and a “No Template control” that contains only H2O or TE buffer in place of template DNA. 3. Initially, it is vital to determine the appropriate cycle number for the reaction (see Note 18). Listed in Table 2 are approximate cycle numbers that will serve as good starting points for cycle number optimization. The cycle number is correct when the “50% control” reaction results in 40–60% of the PCR product of the undiluted control reaction. 4. Specific reaction conditions for two PCR kits are presented below. The first uses the KAPABiosystems LongRange Hot Start PCR kit and has been optimized with human samples. The second used the GoTaq Flexi Kit from Promega and has been optimized for F. grandis samples (see Note 2 for more detailed information). 5. Using the KAPA LongRange Hot Start kit, the reactions are prepared as follows: (a) A master mix is made if several samples are being run simultaneously, which consists of the following components, added in this order: l
24.5μL nuclease-free H2O (for a final volume of 50μL).
l
10μL of 5 buffer solution (vortex at this stage).
l
1μL of BSA in nuclease-free H2O (0.1 mg/mL stock, 2 ng/μL final).
l
1μL of dNTPs (10 mM stock, 200μM final).
l
2.5μL of each primer working solution (10μM stock, 0.5μM final, except O. latipes, 7.5μM stock).
l
3.5μL of MgCl2 (25 mM stock, 1.75 mM final).
l
0.5μL KAPA LongRange Hot Start DNA Polymerase (2.5 U/μL).
l
Vortex and spin.
(b) Aliquot the master mix into the appropriate number of tubes and add 15 ng purified DNA (5μL if diluted to 3 ng/μL) or 5μL of worm lysate as template to the prealiquoted master mix. Also, add 5μL of “50% control” and “No Template control” to the appropriate tubes. (c) The PCR amplification profile is as follows: 94 C for 3 min, followed by the optimized number of cycles of 94 C for 15 s, annealing temperature (Table 2) for 45 s, and 72 C for 45 s. To complete the profile, perform a final extension for 10 min at 72 C.
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6. Using the GoTaq Flexi Kit, the reactions are assembled as follows: (a) A master mix is made if several samples are being run simultaneously, which consists of the following components, added in this order: l
9.5μL nuclease-free water (for a final volume of 25μL).
l
5μL of 5 Colorless GoTaq Flexi Buffer (vortex at this stage).
l
1μL of PCR nucleotide mix (10 mM stock, 400μM final).
l
1μL of each primer solution (10μM stock, 0.4μM final).
l
1μL MgCl2 (25 mM stock, 1 mM final).
l
0.5μL GoTaq DNA polymerase (5 U/μL).
l
Vortex and spin.
(b) Aliquot the master mix into the appropriate number of tubes and add 60 ng purified DNA (6μL if diluted to 10 ng/μL) as template to the prealiquoted master mix. (c) The PCR amplification profile is as follows: 94 C for 2 min, followed by the optimized number of cycles of 94 C for 30 s, annealing temperature (Table 2) for 30 s, and 72 C for 1 min. To complete the profile, perform a final extension for 5 min at 72 C. 7. The primers provided in Table 2 have been tested and verified to result in a single, specific, PCR product; however, when first optimizing the assay, it is recommended to check the specificity of the PCR products on an agarose gel, as conditions may vary slightly based on laboratory equipment and PCR kits used. It is critical to obtain a single product to ensure accurate quantification in step 8. 8. Quantify the resulting PCR products (similar to Subheading 3.2). Add 10μL of each PCR product and 90μL of TE buffer to each of two duplicate wells of a white or black 96-well plate. Also, add 10μL of the DNA concentration standard curve (see Subheading 3.2) and 90μL of TE buffer to each of two duplicate wells of the same plate. 9. Follow steps 6–8 of Subheading 3.2 for fluorescent quantification of DNA. The standards are not used to calculate DNA concentration in this protocol but are useful to assure that the samples fall within the linear range of the instrument and can be helpful when comparing samples on different plates.
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Data Analysis
3.4.1 C. elegans and Human Samples, Real-Time PCR with Standard Curve
1. Obtain mtDNA cycle threshold (CT) values from the RealTime PCR software and average the CT values for the triplicate reactions. If any of the triplicate CT values vary by more than 0.5 units from the others, they should be removed prior to analysis. 2. Obtain the CT values for the standard curve reactions and perform a logarithmic regression with CT on the Y-axis and copy number on the X-axis (see Note 19). 3. Compare the CT values of the samples with those of the standard curve to determine the copy number per PCR reaction. We use the following equation: (a) e[(Sample CT reaction,
slope)/y-intercept]
¼ mtDNA copy number per
4. Assuming worms were picked at a ratio of 1 worm per 15μL and 2μL were used as template, multiply this number by 7.5 to calculate mtDNA CN per worm (see Note 20). 3.4.2 Other Species, Real-Time PCR Without Standard Curve
1. Obtain both mtDNA and nucDNA CT values from Real-Time PCR software and average the CT values from triplicate reactions. 2. To determine the mitochondrial DNA content, relative to nuclear DNA, use the following equations: (a) ΔCT ¼ (nucDNA CT mtDNA CT) (b) Relative mitochondrial DNA content ¼ 2 2ΔCT.
3.4.3 Quantitative, Non-real-Time PCR
1. Obtain the fluorescence values from the plate reader software. 2. Blank subtract—subtract the “No Template Control” values from all other PCR product values (not standard curve values). 3. Average the values of the duplicate wells. 4. Assure that the “50% control” values fall between 40 and 60% of the undiluted control values. 5. The fluorescence values can be used directly to compare relative mitochondrial DNA content between samples.
4
Notes 1. DNA from small-scale worm lysis does not need to be quantified prior to PCR; however, quantification is required for purified DNA from large-scale preparations. 2. The GeneAmp XL PCR kit from Applied Biosystems that was previously used for the QPCR assay has been discontinued. We have now optimized the QPCR assay using KAPA LongRange Hot Start DNA Polymerase with human samples and the
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GoTaq Flexi PCR kit with F. grandis samples. We expect these protocols to be easily adaptable to the other species that we provide primers for. It is also likely that other PCR kits will work with this protocol, though some optimization may be required (discussed in further detail in Subheading 3.4 and Note 17, with extensive examples of troubleshooting steps in Meyer, 2010). 3. Maintaining distinct pre- and post-PCR workstations and pipettes helps to reduce the possibility of contamination of the pre-PCR workstation and, therefore, new PCR reactions, with PCR product. This is especially important when the same PCR target will be amplified repeatedly from many different samples, as is often the case with experiments that use these protocols. We use a PCR hood equipped with a UV sterilizing lamp for reaction assembly, and completed reactions are never opened in this room. This is not a concern with real-time PCR as samples are not processed post-PCR. 4. Six worms in 90μL is the standard condition we routinely use, however, different numbers of worms can be used. For worms at or past the L4 stage, we use a ratio of one worm per 15μL lysis buffer, and for younger worms, we pick one worm per 10μL buffer. 5. We culture worms on K-media plates, as opposed to the standard NGM, because K-media supports thinner bacterial lawns, therefore, resulting in less transfer of bacteria when picking worms for PCR. 6. Samples are picked in triplicate, and each sample is amplified in triplicate, resulting in nine individual real-time PCR reactions per data point. For example, if comparing ethidium bromidetreated worms with controls, we would pick three tubes, each with six worms in 90μL lysis buffer, from the treated group (EtBr 1, 2, and 3) and the same from the control group (C1, 2, and 3). After lysis, each of these samples would be amplified via real-time PCR in triplicate, resulting in nine PCR reactions for each treatment group. 7. When grinding frozen worm pellets, we typically pack the outside of the mortar with dry ice and then chill the mortar and pestle with a small amount of liquid nitrogen. Once the liquid nitrogen has boiled off, the worm pellets are added to the mortar and ground. Care should be taken in the initial few “grinds” as the larger pellets have a tendency to “jump” out of the mortar. 8. We routinely linearize the human mitochondrial DNA from QIAcube preparations by digesting with the PvuII restriction enzyme prior to analysis. This is done not for the short product copy number PCR, but to optimize a long-range PCR for
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DNA damage detection. However, we measure copy number on these linearized templates. While we do not expect this digestion to be necessary for copy number analysis, this has not been exhaustively tested. 9. We prepare larger stocks of the standard curve dilutions that are routinely reused, to reduce variability. Store at 4 C. 10. Depending on the number of worms or cells or the amount of tissue used for DNA extraction, this dilution will change. The aim is to get the DNA concentration in the range of the standard curve for an accurate measurement of concentration. 11. Overhead lab lights are turned off and shades are pulled down. During the 10 min incubation, the plates are covered with aluminum foil or placed in a drawer. 12. PicoGreen working solution should be prepared in excess of what is needed to account for loss during pipetting. We pour the working solution into a reservoir and add 100μL to each well of the plate using an 8-channel micropipette, and typically make between 500μL and 1000μL excess per full 96-well plate. 13. As stated in Note 8, the aim is to get the DNA concentrations in the range of the standard curve. If the DNA concentration is too high, further dilute and measure again. If too low, the undiluted samples can be measured. 14. The standard curve for C. elegans copy number allows us to calculate actual mtDNA CN per worm. Most methods can only determine copy number on a per nuclear copy basis. The standard curve plasmid is pCR2.1 with a 75 bp fragment of the nd-1 gene, containing the real-time PCR primer target sequence, cloned into the multiple cloning site. Based on the molecular weight of the plasmid (MW ¼ 3912) and its concentration, we can accurately determine the number of plasmids (and therefore the number of nd-1 primer targets) per μL. We store this plasmid at a concentration of 10 million copies per μL in single use aliquots (to avoid freeze-thaw cycles) at 80 C. We routinely make duplicate serial dilutions of the standard curve and include both sets of dilutions in our realtime PCR reactions to ensure accuracy. When quantifying purified plasmid DNA, great care needs to be taken to quantify an accurate concentration of DNA, and thereby, plasmid copy number for quantitative use in the standard curve. Initially, a quantification may be taken first by using a NanoDrop or a similar instrument, but then should be more accurately measured by fluorescent PicoGreen quantification as described in Section 3.2. However, it is very important to note that before the PicoGreen quantification, plasmid standards should be linearized with a restriction digest (e.g. for C. elegans
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mitochondrial plasmid, SAC I is a single cutter and can therefore be used to linearize the DNA). Different plasmids and endpoints would need to use a different restriction enzyme to ensure a single cut. The digest should then be column purified using e.g. Qiagen’s PCR Cleanup Kit. If this step is not taken before PicoGreen quantification, the plasmid concentration will be underestimated compared to the true concentration, due to the inability of PicoGreen to completely intercalate into supercoiled plasmid DNA. Additionally, supercoiled plasmid DNA will not amplify properly, also underestimating the actual copy number during quantitative PCR. 15. Individual primers stocks (100μM) are kept at 20 C and working dilutions (25μM) at 4 C. When assembling PCR reactions, working dilutions are mixed 1:1 and further diluted to 5μM each. This primer mix of 2μL is added to each PCR reaction, resulting in final primer concentrations of 400 nM each. 16. As stated in Note 4, each sample is analyzed in three separate real-time PCR reactions, and the results of these three reactions are averaged. 17. In the quantitative, non-real-time PCR for all samples other than C. elegans, the concentration of total DNA is known. Total DNA concentration is based almost entirely on nuclear DNA, and the same amount of total DNA is added to each PCR reaction. For this reason, we effectively start each PCR reaction with the same number of nuclear DNA copies, and the values obtained for relative mitochondrial DNA content do not need to be normalized prior to comparison between samples. 18. PCR product is created proportionally to the amount of specific target sequence in the template during the exponential phase of the reaction. Therefore, for quantitative results, the reaction must be stopped while in this exponential phase. Assuring that the “50% control” sample results in 40–60% of the PCR product of undiluted control will also assure that the reaction is in the exponential phase [12, 13]. 19. When plotting the standard curve CT values, assure that all values fall on the regression line. If a single sample is not on the line, it can be removed. Also, visually inspect the CT values for accuracy. Each time the starting DNA concentration is reduced by half, the CT should increase by 1. 20. If we pick six worms in 90μL of lysis buffer, we assume that 15μL of this lysate is roughly equal to one worm, therefore, copy number per 15μL ¼ copy number per worm. This calculation can be adjusted based on the ratio of worms to volume of lysis buffer.
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Acknowledgments We thank Aleksandra Trifunovic for sharing the mtDNA copy number plasmid for C. elegans with us. This work was supported by R01-ES017540 (JNM), P42ES010356 (JNM), T32-ES021432 (TCL), 1F31ES030588 (TCL), and K99-ES029552 (JHH). References 1. Wallace DC, Chalkia D (2013) Mitochondrial DNA genetics and the heteroplasmy conundrum in evolution and disease. Cold Spring Harb Perspect Biol 5:a021220. https://doi. org/10.1101/cshperspect.a021220 2. Wachsmuth M, Hu¨bner A, Li M et al (2016) Age-related and heteroplasmy-related variation in human mtDNA copy number. PLoS Genet 12:e1005939. https://doi.org/10.1371/jour nal.pgen.1005939 3. Ploumi C, Daskalaki I, Tavernarakis N (2017) Mitochondrial biogenesis and clearance: a balancing act. FEBS J 284:183–195. https://doi. org/10.1111/febs.13820 4. Meyer JN, Leuthner TC, Luz AL (2017) Mitochondrial fusion, fission, and mitochondrial toxicity. Toxicology 391:42–53. https://doi. org/10.1016/j.tox.2017.07.019 5. DeBalsi KL, Hoff KE, Copeland WC (2017) Role of the mitochondrial DNA replication machinery in mitochondrial DNA mutagenesis, aging and age-related diseases. Ageing Res Rev 33:89–104. https://doi.org/10.1016/j. arr.2016.04.006 6. Krasich R, Copeland WC (2017) DNA polymerases in the mitochondria: a critical review of the evidence. Front Biosci (Landmark Ed) 22:692–709 7. Campbell CT, Kolesar JE, Kaufman BA (2012) Mitochondrial transcription factor A regulates mitochondrial transcription initiation, DNA packaging, and genome copy number. Biochim Biophys Acta 1819(9–10):921–929 8. Moraes CT (2001) What regulates mitochondrial DNA copy number in animal cells? Trends Genet 17:199–205. https://doi.org/10. 1016/S0168-9525(01)02238-7 9. Dorn GW, Vega RB, Kelly DP (2015) Mitochondrial biogenesis and dynamics in the developing and diseased heart. Genes Dev 29:1981–1991. https://doi.org/10.1101/ gad.269894.115 10. Pickles S, Vigie´ P, Youle RJ (2018) Mitophagy and quality control mechanisms in mitochondrial maintenance. Curr Biol 28:R170–R185. https://doi.org/10.1016/J.CUB.2018.01.004
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Chapter 9 Methods to Monitor Mitophagy and Mitochondrial Quality: Implications in Cancer, Neurodegeneration, and Cardiovascular Diseases Simone Patergnani, Massimo Bonora, Esmaa Bouhamida, Alberto Danese, Saverio Marchi, Giampaolo Morciano, Maurizio Previati, Gaia Pedriali, Alessandro Rimessi, Gabriele Anania, Carlotta Giorgi, and Paolo Pinton Abstract Mitochondria are dynamic organelles that participate in a broad array of molecular functions within the cell. They are responsible for maintaining the appropriate energetic levels and control the cellular homeostasis throughout the generation of intermediary metabolites. Preserving a healthy and functional mitochondrial population is of fundamental importance throughout the life of the cells under pathophysiological conditions. Hence, cells have evolved fine-tuned mechanisms of quality control that help to preserve the right amount of functional mitochondria to meet the demand of the cell. The specific recycling of mitochondria by autophagy, termed mitophagy, represents the primary contributor to mitochondrial quality control. During this process, damaged or unnecessary mitochondria are recognized and selectively degraded. In the past few years, the knowledge in mitophagy has seen rapid progress, and a growing body of evidence confirms that mitophagy holds a central role in controlling cellular functions and the progression of various human diseases. In this chapter, we will discuss the pathophysiological roles of mitophagy and provide a general overview of the current methods used to monitor and quantify mitophagy. We will also outline the main established approaches to investigate the mitochondrial function, metabolism, morphology, and protein damage. Key words Metabolism, Mitochondrial morphology, Pathology, Mitochondrial quality control, Cardiovascular diseases (CVD), Homeostasis
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Introduction The functions attributed to the mitochondria among the more general cellular activities have been progressively increasing during the last decades, strongly changing the textbook image of an intracellular organelle, principally seen as an energy provider. A more precise draw has to consider mitochondria as pivotal regulators of
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cell death, being it able to respond to different death stimuli through the release of mitochondrial proteins able to trigger the proapoptotic routine. Presence of specialized zones, where mitochondria contact the endoplasmic reticulum (ER), allows for a regulated exchange of lipids, cholesterol, and in particular, Calcium (Ca2+) [1–4], whose influx and outflow influence both mitochondrial enzymatic activities or specialized cytoplasmic functions like muscle contraction and others. The outer mitochondrial membrane (OMM) behaves as a dock surface for several proteins involved in the inflammatory and antiviral response [5], and, in addition, the release of mitochondrial(mt)DNA and formyl peptides from mitochondria, both intra or extracellularly, strongly prompts the inflammatory response. Mitochondria are not only providers of adenosine triphosphate (ATP), but also generators of metabolic intermediates to be used for the synthesis of fatty acids and amino acids and are so flexible to switch among these different functions to meet the cellular requirements. Mitochondria are also the source of iron-sulfur clusters, which are present in a wide number of mitochondrial and not mitochondrial proteins [6]. Being mitochondria as regulatory platforms of different cellular functions is not surprising that not only their specific functions, but also their number and the quality of their activities have to be strictly regulated. Mitochondria usually are present in the cells under the form of a dynamic network, where mitochondrial mass increases as a consequence of mitochondrial biogenesis, while its diminution can occur through a selective form of autophagy, termed mitophagy. Autophagy is a more general mechanism that is devised to destroy and recycle intracellular organelles, parts of cytoplasm and unfolded or aggregated proteins, lipid droplets, and xenobiotics. Although being important to recycle metabolic intermediate and fuel cell metabolism under shortage conditions, the autophagic process itself is something more than a mere degradative process. Instead, it is devised to reshape the cellular composition, often eliminating, selectively, the damaged or nonfunctional part of the cell. Autophagy is under the control of the regulatory kinases mechanistic target of rapamycin (mTOR) and 50 adenosine monophosphate-activated protein (AMPK). mTOR exerts a general inhibitory drive on autophagy, and its activity is reduced when the overall metabolic demand exceeds the supply. On the contrary, AMPK promotes autophagy, and it can be activated by impairment of mitochondrial activity, in terms of lesser ATP synthesis, reduced oxygen (O2) consumption, increased reactive oxygen species (ROS) production, or Ca2+ mobilization [7]. Mitophagy regulation is not a completely clarified topic yet. During short-term starvation, mitochondrial pool is not depleted, to not further reduce cellular production of energy, while general autophagy can occur to sustain oxidative metabolism [8]. Prolonged starvation can lead to mitochondrial depletion through a nonspecific mechanism that can also involve other organelles and
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cytoplasm. On the contrary, mitochondrial ROS production or depolarization can trigger mitophagy machinery, which is organized into different moments: the identification of the parts of the mitochondrial network to address to demolition and the recruitment to the autophagic machinery, which terminates with fusion to lysosomes. There are several mechanisms by which mitophagy is induced. The first observation concerned reticulocytes that, during differentiation, loss mitochondria by mitophagy throughout the activity of NIP3-like protein X (NIX/BNIP3L). Another mechanism was found upon hypoxic conditions, where the OMM protein FUNDC1 works as a mitophagic receptors, thanks to its microtubule-associated proteins 1A/1B light chain 3B (LC3)binding domain at N-terminal level [9]. Similarly, FK506-binding protein 8 (FKBP8) and BCL2L13 were identified as a LC3-interacting proteins capable to activate mitophagy [10, 11]. Undoubtedly, the most studied is the molecular mechanism regulated by the PTEN-induced kinase 1 (PINK1)/Parkin axis. PINK1 and Parkin belong to a series of genes referred to as PARK genes. Mutations in these genes (α-synuclein (α-syn, PARK1/4), parkin (PARK2), PINK1 (PTEN-Induced Kinase 1— PARK6), DJ-1 (PARK7), LRRK2 (PARK8), and ATP13A2 (PARK9)) have been linked to familiar forms of Parkinson’s disease (PD). Studies occurring in these PD forms have permitted to provide insight into the molecular mechanisms of mitophagy. In particular, it has been demonstrated that PINK1 and Parkin act in a common pathway to regulate mitochondrial function, autophagy, and protein accumulation. From its discovery in 1998, about 100 mutations have been identified for Parkin gene as a causative gene for autosomal recessive parkinsonism [12]. The Parkin gene product is a cytosolic protein characterized by an ubiquitin-like (UBL) domain at the N-terminus. Originally linked to autosomal recessive early-onset PD in 2004, PINK1 encodes a ubiquitous protein characterized by a mitochondrial targeting sequence (MTS), a transmembrane domain, and a highly conserved serine/threonine kinase domain. At today, about 30 pathogenic PINK1 mutations have been identified to impair its kinase activity and provoke loss of function [13– 15]. Normally, the levels of PINK1 are very low. This is due to a fine-tuned mechanism that begins with the import of PINK1 into mitochondria by the activity of the translocases of the inner (TIM23) and outer (TOM) membrane complex (Fig. 1). Once arrived in the inner mitochondrial membrane (IMM), PINK1 is subjected to series of proteolytic cleavages that reduce the fulllength form of PINK1 into fragments, which are then degraded by proteasome [16–18]. When stress conditions increase and mitochondria suffer damages, the activity of TIM23/TOM complex is disrupted, and PINK1 accumulates on the OMM. Here, once stabilized by a molecular complex composed of TOM proteins
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Fig. 1 Schematic representation of the molecular process of mitophagy. During resting conditions, PINK1 is imported into IMM via TIM-23-TOM activity. Here, PINK is cleaved by PARL in fragments, which are then degraded by proteasome. Upon damage, import of PINK1 is blocked leading to accumulation and stabilization in OMM surface, thanks to TOM proteins. In this conformation, PINK1 mediates phosphorylation events aimed to activate and recruit Parkin at OMM. Once activated, Parkin can elongate Ub chains, and a series of autophagic receptors (NDP52, OPTN, NBR1, TBK1, and p62) are recruited. Now, the damaged mitochondria can be surrounded by autophagosomal membrane to form the mitophagosome
[19, 20], PINK1 phosphorylates itself at S402, S228, and T257 and converts the autoinhibited E3-ubiquitin (Ub) ligase Parkin into an active phospho-Ub-dependent enzyme [21, 22]. In this active state, Parkin actively ubiquitinates several mitochondrial proteins at the OMM. Ubiquitination events promote the recruitment of the Ub-binding autophagy receptors p62/Sequestome, NBR1, NDP52, optineurin (OPTN), and TAX1BP1 (TBK1), which connect the damaged mitochondria to phagosomes for clearance in the lysosome (Fig. 1) [23–25]. 1.1 Role of Mitophagy in Pathology
Mitochondria dysfunction and mitophagy defects are broadly implicated in numerous human diseases, including cardiovascular diseases (CVD), cancers, and neurodegenerative diseases. Below, we discuss contributions of mitophagy in pathologies, which are widely emergent in recent years.
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Mitophagy is essential for cardiovascular homeostasis and for cardiac mitochondrial quality control, as well as for the maintenance of the cardiac heart under both normal and pathophysiological conditions [26–29]. Aberrant mitophagy contributes to activate matrix metallopeptidase -9 (MMP-9), causing degradation of the important gap junction protein connexin-43 (Cx-43) in the ventricular myocardium. Reduction in Cx-43 levels was associated with increased fibrosis and ventricular dysfunction in heart failure. Recent insight by Tong M. and colleagues has shown that the suppression of mitophagy increased the accumulation of lipids in the heart during high-fat diet consumption. The same study revealed that the transcriptional activator (TAT)-Beclin1, which enhances mitophagy, tends to inhibit the development of diabetic cardiomyopathy [27]. Notably, PINK1 and Parkin protein levels are significantly decreased in type 1 diabetic hearts, resulting in decreased cardiac autophagy, which was correlated with reduced expression of the small GTPase Rab9, responsible for mitochondrial degradation during erythrocyte maturation in the absence of ATG5. Increased level of Rab9 protein was higher in mitochondria deficiency, suggesting that Rab9 mediated degradation of mitochondria in diabetic heart and contributed to mitophagy under certain conditions [30–32]. In recent investigated work, it has been found that sirtuin 1 (SIRT1), a deacetylase stimulated in response to several cardiac stresses to promote cell survival, promoted the induction of mitophagy in response to ER stress and eventually protected the cardiac cells from cell death. However, the inhibition of SIRT1 decreased ER stress-induced mitophagy. The whole-body deletion of PGAM family member 5 (PGAM5), a mitochondrial serine/threonine protein phosphatase, activates PINK1-mediated mitophagy in the heart. Moreover, PGAM 5 increased infarct size in mice with cardiac ischemia/reperfusion, which is implicated in mitophagy inhibition in the myocardium [33]. Hoshino and associates found that nuclear p53 attenuates mitophagy in the ischemic heart, which causes accumulation of altered mitochondria and increased myocardial damage [34]. The mitochondrial fission dynamin-1-like protein (DRP1), which is a critical fission regulatory, has an important role in the heart mitophagy. DRP1 deletion provokes mitophagic inhibition and consequent cardiac damage and enhanced risk to myocardial ischemia-reperfusion injury [35, 36]. Furthermore, DRP1 mutation C452F causes spontaneous development of monogenic-dilated cardiomyopathy with abnormal mitochondrial morphology and defective mitophagy [37]. Contrasting researches have revealed that mice lacking mitochondrial fusion or fission proteins manifest deteriorations in the developmental cardiac and increased susceptibility to cardiac injury [38, 39]. The ratio of mitofusin 2 (MFN2; a fusion protein) and
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DRP1 (a fission protein) was decreased during heart failure, suggesting increased mitophagy [40]. Nevertheless, loss of the mitochondrial division DRP1and Parkin activates the mitophagic aberration and subsequently reduces mitochondrial respiration and provokes lethal cardiac aberrations. 1.1.2 Mitophagy in Cancer
Mitophagy prevents oncogenic transformation and maintains cellular homeostasis and eventually can be considered as a tumor suppressor process because selectively degrading impaired mitochondria can prevent the accumulation of ROS [1, 41]. The repression of mitophagy leads to an accumulation of impaired mitochondria and subsequently enhances tumorigenesis. Increased evidence suggested the functional loss of mitophagy regulators in the development and the progression of cancer, and this became evident through the modulation of expression of the regulators including Parkin, BNIP3, NIX, and representative of tumor suppressor such as Rb, p53, and oncogenes like NF-kB, Hif-1a, and FOXO3 [42]. Interestingly, Parkin is a most frequently deleted gene in cancers; its deletion is associated with various types of cancer including serous ovarian breast carcinomas, liver, and colon cancer [43]. Besides its role in mitophagy, Parkin can behave as a tumor suppressor modulating the levels of various key cell cycle proteins, such as Cyclin D1, Cyclin E, and CDK4, all having a role in controlling G1/S progression. Further studies supported the notion that Parkin acts as a tumor suppressor: they knocked down Parkin and thus showed an increase in spontaneous liver tumors and sensitized mice to γ-irradiation-induced tumorigenesis [44]. Several insights showed that the dysregulation of proteins involved in the mitophagic process such as the promitophagic receptor BNIP3 has a tumor promoter role in melanoma, renal cell carcinoma, and pancreatic cancer, whereas in breast cancer, tumor suppressor functions [45]. Furthermore, knocking down BNIP3 stimulates tumor formation and metastasis in mouse model, suggesting a key role of mitophagy in suppressing cancer development [42]. Again, BNIP3 is frequently been deleted in triple-negative breast cancer (TNBC) and epigenetically silenced in other types of cancer including gastric, lung, liver, and pancreatic cancer [42]. Hypoxia occurs in solid tumor. It has been found that HIF-1α plays a crucial role in adaptation of cancer cells under hypoxic condition, and it enhances the expression of BNIP3. Furthermore, an enhanced expression of HIF-1α has been found in hemangioblastoma, glioblastoma multiforme, breast and prostate cancer, colonic adenocarcinoma, and subtypes of the lung [46].
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Mitophagy plays a protective role in numerous neurodegenerative diseases via mitophagy by clearing impaired mitochondria in neuronal axon, and genetic aberrations in both PINK and Parkin have been associated with the development of hereditary PD [47]. Mutations in α-synuclein (α-syn), a component of Lewy bodies, the pathological hallmark of PD, leads to a block in mitophagy [48]. Overexpression of A53T α-syn mutant leads to a stimulation of p38 MAPK and thus phosphorylated Parkin at SER131 to disrupt its functions [49]. Recently, the activity of α-syn has been correlated to the activity of mitochondrial Rho GTPase 1 (Miro1). This protein anchors mitochondria to microtubule motors and is required for mitochondrial motility in healthy conditions to stop mitochondrial motility and initiate mitophagy, Miro1 should be removed. On the contrary, α-syn interacts with Miro1 via its N-terminus and enhances Miro1 protein levels, thus leading to an abnormal Miro1 accumulation on the mitochondrial surface and delayed mitophagy [50]. Other genes and proteins that can mediate between mitochondrial autophagy and the progression of PD. The protein DJ1, associated with a rare form of autosomal recessive PD, is localized to mitochondria, where it regulates the removal of endogenous ROS and the mitophagy process [51, 52]. Deletion of DJ1 enhanced the recruitment of Parkin to damaged mitochondria and increased mitochondrial autophagy [53]. Leucine-rich repeat kinase 2 (LRRK2) is present in the mitochondria, modulates mitochondrial homeostasis, and represents the major risk factor for PD and it having been shown to be correlated with sporadic and hereditary PD. G2019S, one of the most frequent LRRK2 mutations implicated in PD, leads to fission of mitochondria via an increased DRP1 activity, followed by activation of mitophagy [54]. In addition, dysfunctional mitochondria are found in diseases of the peripheral nerves, including the inherited axonal neuropathies, Charcot-Marie-Tooth neuropathy disease type 2 (CMT2A) is commonly associated with aberrations in proteins linked to mitochondrial dynamic including MFN2 and GDAP1 [55]. A recent work showed that motor neuron of CMT2A patients and expressing mutant MFN2 exhibit defects in mitophagy. These findings highlight the importance of mitophagy process to axonal homeostasis [56]. Parkin is also associated with the progression of Alzheimer’s disease (AD) [57]. Human wild-type full-length Tau (hTau) affects the mitochondrial membrane, thereby causes mitochondrial dysfunction. A recent evidence found that both hTau and familial forms of frontotemporal dementia (FTD) mutant Tau (hP301L) suppressed mitophagy in neuroblastoma cells via the reduction of Parkin. The hTau NH2-terminal fragment modulates Parkin via the repression of ANT-1-dependent ADP/ATP exchange, inhibits mitophagy, and affects synaptic degeneration in AD [58].
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Sirtuins family are NAD+-dependent enzymes disrupted in AD and contributing to AD pathogenesis. Increased evidence reported that SIRT1 modulates multiple pathways that go amiss in AD, including neuroinflammation, neurodegeneration, and mitochondrial alterations [59]. Resveratrol has been shown to enhance the expression of SIRT1 and can upregulate the level of LC3-II/LC3I, Beclin 1, and Parkin, reduce the positioning of LC3 and TOM20, as well as regulate BNIP3- and NIX-related pathways in AD-excessive SIRT2 stimulation leading to mitochondrial alteration and ensuing mitophagy [60]. Alteration in mitophagy is linked also to Huntington’s disease (HD). Dysfunctional mitochondria are correlated with different animal models of HD, and mutant huntingtin (mhtt) influences mitochondrial biogenesis and function and affects negatively the mitochondrial delivery to the lysosome [61]. In addition, valosincontaining protein (VCP), known as p97, is located in various subcellular organelles, such as ER, nucleus, and mitochondria, where it has different functions including regulation of ER and mitochondria degradation, autophagy, and DNA repair [62]. An elegant work has shown that VCP selectively translocates to the mitochondria and acts as an mtHtt-binding protein on the mitochondria. Accumulated in mitochondria, VCP elicits excessive activation of mitophagy, inducing neuronal cell death. By adding the HV-3 peptide, VCP translocation to the mitochondria is abolished, the excessive mitophagy is inhibited, determining a reduction of cell death in both HD mouse/patient-derived cells and HD transgenic mouse brains [63]. The α-tubulin deacetylase HDAC6 is a cytosolic histone deacetylase suggested to block mitophagy. Thus, inhibiting HDAC6 increases neuronal vesicular flux and is considered as a potential therapeutic to treat HD, but the mechanisms remain unclear [64, 65]. Several evidence in cell lines suggest the role of HDAC6 for autophagosome–lysosome fusion and also that HDAC6 promotes the removal of mHtt and impaired mitochondria [66, 67]. In amyotrophic lateral sclerosis (ALS), mitochondria are damaged and accompanied with the depletion of motor neuron disease (MND) and upper motor neuron (UMN). Increased evidence suggest the potential role of mitophagy in ALS. More than 30 gene mutations including OPTN, TBK1, VCP, C9ORF72, and SOD1 are linked to ALS. Consistently, loss of function mutations in genes TBK1 and OPTN are involved in the impairment of mitophagy pathway and accumulation of damaged mitochondria [68]. Approximately, 20% of the ALS are caused by aberration of SOD1, which was the first ALS gene discovered [69]. For instance, mutant SOD1 causes mitochondrial dysfunction and contributes to motor neuron demise in ALS, thus stimulating mitochondria
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quality control (MQC) to reduce impaired mitochondria accumulation via mitophagy process [70, 71]. It has been reported that SOD1 relies on PINK1 to decrease the mitochondrial dynamic protein Miro1. Mutant SOD1 altered axonal transport in PINK1/Parkin pathway-dependent manner [72].
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Monitoring Mitophagy Using Fluorescent Probes To determine the colocalization of mitochondria with either autophagosomes or lysosomes, represent the main method to measure mitophagy [73]. The colocalization of mitochondria with autophagosome can be studied by using a green fluorescent protein (GFP)LC3 plasmid (for autophagosome labeling) and a mitochondrially targeted red fluorescent protein (RFP) (to visualize mitochondria) [74]. Alternatively, it is possible to monitor the delivery of sequestered mitochondria to the lysosome by using the fluorescent dyes MitoTracker and LysoTracker [75] or the mitochondrial pH-dependent Keima protein (named mito-Keima). This fluorescent protein changes color depending on the pH environment, and when mitochondria undergoing mitophagy are sequestered into the lysosomal compartment, the peak of excitation of mito-Keima shifts from green (neutral pH) to red (acidic) color [76, 77]. Following a PINK1 stabilization, Parkin protein translocates from cytosol to mitochondria. Fluorescent recombinant chimeras of Parkin, such as mCherry-Parkin, permit to monitor this translocation and thus understand when the mitophagic process is activated at OMM surface. The same information may be obtained performing an immunofluorescence assay against Parkin and a mitochondrial marker [78, 79]. In the following sections, we will describe two different assays for monitoring and quantifying the mitophagic levels by using a fluorescence approach.
2.1 Colocalization Autophagosome and Mitochondria by Confocal Fluorescent Microscopy
1. Biological material (cell cultures of interest) (see Note 1).
2.1.1 Materials
3. Glass coverslips 24 mm in diameter (see Note 2).
2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, and 1% L-glutamine. 4. Six-well culture plates. 5. 1 phosphate-buffered saline (PBS) buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 6. 10 mM carbonyl 4-(trifluoromethoxy)phenylhydrazone (FCCP).
cyanide
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7. Plasmid encoding for a fluorescent mitochondrial probe, that is, mCherry-mito7 [80] mt-DsRed [81] (see Note 3). 8. DNA transfection reagent, that is, jetPEI® (PolyPlus transfection, 101-10 N). 9. Attofluor cell chamber (e.g., ThermoFisher, A7816). 10. 4% paraformaldehyde (PFA) solution, pH 7.4, in 1 PBS. Add 4 g of PFA in 100 ml of 1 PBS. 11. Inverted Zeiss LSM 510 confocal microscope. 2.1.2 Methods
1. Plate cells onto coverslips (24 mm in diameter) at a density of 100,000 cells per well in a 6-well plate. 2. Let the cells grow on the cover slip surface to achieve 60–75% confluence. 3. Transfect cells with 1 μg/coverslips GFP-LC3 plasmid and mCherry-mito7 by using the appropriate method. 4. After 4–6 h, replace the transfection medium with warm medium culture. 5. Wait for 24–48 h (see Note 4). 6. Add a final concentration 1 μM of FCCP to provoke mitophagy (see Note 5). 7. After 3 h, image the cells. Alternatively, cells can be fixed in 4% PFA 15 min at room temperature (RT), washed three times with 1 PBS, and stored at 4 C in the dark (see Note 6). 8. Move the coverslips into a cell chamber and place in a 37 C thermo-controlled stage of the inverted Zeiss LSM 510 confocal microscope (see Note 7). Fluorescence images are captured providing excitation at 488 nm (green) and 546 nm (red) with detection of green (500–540 nm) and red (580–640 nm) fluorescence signals. 9. To obtain a statistical measurement, acquire at least 20–25 images per condition (see Note 8).
2.1.3 Data Handling
Image processing may be achieved by using the microscope software (ZEN elements for Zeiss instruments) or Fiji software. After background subtraction on both green and red channels, generate merged images to visualize the colocalization amounts (Fig. 2). To quantify the yellow signal, we suggest using the colocalization plug-in available in Fiji software or the colocalization command of ZEN element software. Alternatively, it is possible to perform a simple count of yellow dots or measure the intensity profile of a region of interest (ROI) for both red and green channels (see Note 9).
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Fig. 2 Colocalization of the autophagosomal marker GFP-LC3 and mitochondria. After transfection, cells were treated with FCCP to cause mitochondrial damage and the accumulation of autophagosome. LC3 (green) and mitochondrial (red) signals were then recorded by using a fluorescence confocal microscope equipped using an oil 63 1.4 NA lens, and the colocalization rate was investigated by merging the channels. The regions indicated in blue box are enlarged in the Zoom panel. Yellow dots are representative of colocalization of GFPLC3 and mCherry-mito7. Scale bar 10 μm 2.1.4 Notes
1. It is also possible to use cells with stable expression of GFPLC3 construct. However, the establishment of a stable cell line requires a significant investment of time. In addition to this, stable cell lines are often immortalized tumor cell lines and are far from the physiology of primary cells. 2. Alternative supports are cell imaging dishes 35 mm in diameter. 3. To stain mitochondrial structures, a large series of mitochondrial-specific fluorescent dyes (MitoTracker Green, MitoTracker Orange, MitoTracker Red, and MitoTracker Deep Red) may be used.
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4. Certain transfection protocols may induce autophagy. We recommend leaving the cells for a period of 48 h posttransfection to allow equilibration [82]. 5. Adjust the FCCP concentration according to the cell type used. 6. Fixation procedure may produce autofluorescent puncta or a reduction of GFP-LC3 staining. 7. Changes in temperature and prolonged light exposure during live-cell imaging may provoke photodamage and photobleaching. Keep laser potency and exposure time low. 8. Keep exposure time and illumination constant across samples. 9. The overlay of the channels of interest possesses some limitations. The merged staining may be dependent on the signal intensity of each channel. To avoid artifacts, we suggest verifying the result obtained by using algorithms performing intensity correlation coefficient-based analyses (ICCB), such as Pearson’s or Manders’ coefficients and object-based approaches (such as the centroid or intensity center calculation). 2.2 Colocalization Parkin and Mitochondrial Marker
1. Biological material (cell cultures of interest).
2.2.1 Materials
3. Glass coverslips 24 mm in diameter (see Note 1).
2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 4. Six-well culture plates. 5. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 6. Permeabilization Buffer: 0.05–0.3% Triton X-100 (or 100 μM digitonin or 0.5% saponin) in 1 PBS. 7. Blocking Buffer (chose one of below): – 5–10% serum from host species of secondary antibody (blocking). – 1–3% bovine serum albumin (BSA) in 1 PBS. 8. 4% PFA solution, pH 7.4, in 1 PBS. Add 4 g of PFA in 100 ml of 1 PBS. 9. Primary antibody mouse anti-ATP5A (abcam, ab14748) (or another mitochondrial marker). 10. Primary antibody PA5–13398).
rabbit
anti-Parkin
(ThermoFisher,
11. Secondary antibody Alexa Fluor 488 goat anti-rabbit (ThermoFisher, A27034).
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12. Secondary antibody Alexa Fluor 647 goat anti-mouse (ThermoFisher, A28181). 13. Antigen Retrieval Buffer (ARB): 100 mM Tris, 5% (w/v) urea, pH 9.5. 14. Inverted Nikon confocal microscope system A1 R (see Note 1). 2.2.2 Methods
Sample Preparation and Transfection 1. Plate cells onto coverslips (24 mm in diameter) at a density of 100,000 cells in a 6-well plate. 2. Wait at least for 24 h. 3. After three washes with 1 PBS, fix cells in 4% PFA 15 min at RT. 4. Wash three times with 1 PBS (see Notes 2 and 3). 5. Add 1 ml of permeabilization buffer in each well. Incubate for 10 min at RT with agitation. 6. Wash three times with 1 PBS. 7. Incubate cells with 1 ml of blocking buffer for 30 min at RT with agitation to block unspecific binding of the antibodies (see Note 4). 8. Incubate cells in the diluted antibody in 1% BSA in 1 PBS in a humidified chamber for 1 h at RT overnight (ON) at 4 C (see Note 4). 9. Wash three times with 1 PBS. 10. Incubate cells with the secondary antibodies in 1% BSA in 1 PBS for 1 h at RT in the dark. We recommend a dilution range of 1:500–1:1000. 11. Wash three times with 1 PBS. 12. Mount coverslips with a drop of mounting medium (see Note 5). 13. Store in the dark at 4 C or 20 C. Measurement 14. Move the coverslips in a controlled stage of the inverted microscope (see Note 3). Fluorescence images are captured, providing excitation at 488 nm (green) and 647 nm (deep red) with detection of green (500–540 nm) and red (650–720 nm) fluorescence signals (see Note 4). 15. To obtain a statistical measurement of the mitochondrial network, acquire at least 20–25 images per condition (see Note 5).
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2.2.3 Data Handling
Image processing may be achieved by using the microscope software (NIS elements for Nikon instruments) or Fiji software. After background subtraction on both green and red channels, generate merged images to visualize the colocalization amounts. To quantify the yellow signal, we suggest using the colocalization plug-in available in Fiji software or the colocalization command of NIS-element software (Fig. 3). Alternatively, it is possible to perform a simple count of yellow dots or measure the intensity profile of a ROI for both red and green channels.
Fig. 3 Colocalization Parkin and mitochondrial marker. Cells were fixed, permeabilized, and exposed to primary antibodies against ATP5A (mitochondrial marker, red) and Parkin (green). After incubation with secondary antibodies, cells were imaged in a confocal inverted microscope, and the colocalization rate was investigated. Regions in blue box are enlarged in the Zoom panels. Yellow dots are representative of colocalization of Parkin and mitochondria. Scale bar 10 μm
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1. Alternative supports are glass coverslips 13 mm in diameter or cell imaging dishes 35 mm in diameter. 2. Before permeabilization, it is possible to perform the antigen retrieval by preheating the coverslips with the ARB at 95 C. After 10 min, remove the coverslips and wash cells in PBS three times for 5 min. 3. Proceed with permeabilization or keep sample immerged in PBS at 4 C for 1 week. 4. Blocking and primary antibody buffers may be supplemented with 0.01% Triton X-100 (or 10 μM digitonin or 0.1% saponin). 5. We recommend sealing coverslips with nail polish to prevent drying.
3
Monitoring Mitophagy Using Nonfluorescence Methods Undoubtedly, fluorescence methods are the most used techniques to study mitophagy. However, these assays only demonstrate colocalization of the autophagosome with mitochondria. They cannot be used to determine mitochondrial degradation. Furthermore, the use of fluorescent indicators may have several limitations. For example, MitoTracker dyes are dependents of mitochondrial potential. Changes in lysosomal pH can affect the staining pattern of Lyso tracker. GFP-LC3 puncta are not always autophagosomes and may form aggregates that give misleading results. Clearly, other approaches measuring mitophagy should be combined with these assays to confirm the results obtained. Transmission electron microscopy (TEM) is one of the most powerful approaches for the evaluation of mitophagy. Fulfilling the philosophy of “seeing is believing,” the principle of this analysis is the direct identification of autophagosomes and autophagolysosomes that have physically engulfed a mitochondrial particle, by visual inspection. Indeed, the high resolutive power of TEM allows recognition of the morphological features, which characterize autophagosomes, autophagolysosomes, and mitochondria. It should be noted that mitophagy evaluation by TEM could sometimes be problematic. This is mainly due to: (1) poor sampling of the phenomenon (depending on the cell orientation, during cutting autophagosomes might appear in the section or not), (2) methodological artifacts deriving from fixation, which might lead to misinterpretation, and (3) a specialized training to operate at is required. Considering these remarks, complementary techniques should be considered. Following, we will describe two methods determining the mitochondrial autophagy by using immunoblot and enzyme-linked immunosorbent assay (ELISA) procedures.
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3.1 Evaluation of Mitochondrial Protein Amounts
3.1.1 Materials
The final step of mitophagy is the removal of the sequestered mitochondria [24]. Western blotting can be used to measure mitochondrial protein degradation to determine whether mitophagic process is activated. When this experiment is conducted, all mitochondrial compartment proteins (including matrix proteins) should be analyzed. The only detection of outer mitochondrial membrane proteins, described to be mitophagic targets (TOM20, VDACs, and mitofusins), can mislead the result because these proteins are also degraded by the proteasome. 1. Biological material (cell cultures of interest). 2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 3. FCCP (see Note 1). 4. Radioimmunoprecipitation (RIPA) lysis buffer (150 mM sodium chloride, 1.0% NP-40 or Triton X-100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 50 mM Tris, pH 8.0, supplemented of protease and phosphatase inhibitors). 5. Transfer Buffer 1 (1 l): Tris base 5.8 g, glycine 2.9 g, SDS 0.37 g. Make 800 ml with dH2O, then add 200 ml MeOH. 6. Tris-Buffered Saline (TBS; 10): 1.5 M NaCl, 0.1 M Tris– HCl, pH 7.4. 7. TBS containing 0.05% Tween-20 (TBS-T). 8. Blocking Solution: 5% milk in TBS-T. 9. Precast gel 4–12% NW04120BOX).
Bis-Tris
Bolt
(ThermoFisher,
10. Nitrocellulose or PVDF membranes. 11. Primary antibody anti-HSP60 (mitochondrial matrix marker, mouse, Santa Cruz, sc-13115), anti-TIM23 (inner mitochondrial membrane marker, mouse, BD Bioscience, 611222), and anti-pan/VDAC1 (outer mitochondrial membrane marker, rabbit, abcam, ab34726) (see Note 2). 12. Primary antibody anti-GAPDH (loading marker, rabbit cell signaling, 5174). 13. Primary antibody anti-PDI (ER marker, rabbit, cell signaling, 2446) (see Note 3). 14. Restore western blot stripping buffer (ThermoFisher, 21059). 3.1.2 Method
Sample Preparation and Homogenates Collection 1. Plate cells in 6-well plates at a density of 150,000 cells. 2. After at least 24 h, treat cells with a strong mitophagic inducer, such as 100 nM FCCP.
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3. Collect cellular homogenates and lyse them with RIPA lysis buffer. Keep homogenates in ice. After 30 min, centrifuge cell lysates at 12,000 g for 10 min at 4 C, collect the supernatant, and keep them at 80 C. Western Blot. After having determined the protein concentration for the cell lysate, prepare the sample (10–15 μg of proteins) in denaturating conditions and boil the mixture at 95–100 C for 5 min. 4. Load samples in a precast gel 4–12% and perform the electrophoresis at 150 mV for 30–45 min. 5. Following electrophoresis, activate PVDF with methanol for 1 min and rinse with transfer buffer. If a nitrocellulose membrane is used, only rinse it with transfer buffer. Assemble the transfer stack with sponges, filter papers, membrane, and gel, being careful to that the gel is facing the cathode () and the membrane side is facing the anode (+). We recommend transferring the gel for 2 h at 350 mA. 6. After the protein transfer, rinse the membrane in TBS-T and place the membrane in blocking solution and incubate with agitation for 1 h. 7. Place the blot in the primary antibodies solution and incubate with agitation overnight. Wash the blot three times for 5 min. 8. Place the blot in the secondary antibody solution and incubate with agitation for 1 h at RT. Wash the blot three times for 5 min. 9. Proceed with chemiluminescent detection. In the case some proteins have similar molecular weight, stripping and reprobing the blot as follows: 10. After detection, wash blot to remove chemiluminescent substrate. 11. Incubate blot in Restore western blot stripping buffer for 15 min at RT. Wash the blot three times in TBS-T with agitation for 10 min at RT. 12. Place the membrane in blocking solution and incubate with agitation for 1 h. 13. Place the blot in the primary antibodies solution and incubate with agitation O.N. Wash the blot three times for 5 min. 14. Place the blot in the secondary antibody solution and incubate with agitation for 1 h at RT. Wash the blot three times in TBS-T with agitation for 10 min at RT. 15. Proceed with chemiluminescent detection.
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Fig. 4 Evaluation of mitochondrial protein amounts. Levels of mitochondrial proteins were analyzed by western blot. Primary antibodies against HSP60 (matrix marker), pan/VDAC1 (OMM marker), and TIM23 (IMM marker) were used, and the specific levels measured were normalized with GAPDH. PDI (ER marker) was used as control
3.1.3 Data Handling/ Processing
3.1.4 Notes
Quantify the bands with a densitometric program and then divide the protein amounts of each mitochondrial marker with the loading marker. 1. To detect mitochondrial protein degradation is necessary to have high levels of mitophagy. We suggest using this technique following treatment with a mitochondrial stressor inducer like FCCP. 2. Considering that is not clear whether mitophagy is the only cellular mechanism to remove mitochondria, it is recommended to verify the protein amounts of all mitochondrial subcompartments. 3. To ensure that the protein loss is limited to the mitochondrial compartment, it is recommended to verify the protein amounts of another intracellular compartment (e.g., ER proteins) (Fig. 4).
3.2 Parkin Translocation by Subcellular Fractionation
Mitophagy is a homeostatic process that targets unwanted and altered mitochondria to autophagosomes with subsequent degradation in autolysosomes. The selectivity of mitophagic response is achieved through specific mitophagic receptors (such as NDP52 and OPTN), which recognize cargos tagged with degradation signals as well as the autophagosomal membrane protein LC3 [25, 83, 84]. NDP52 and OPTN are cytosolic receptors recruited by PINK1 in the first steps of mitophagy. They are responsible to
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recognize the damaged mitochondria and to promote its sequestration into phagosome. To potentiate the mitophagic response, PINK1 phosphorylates Parkin, an E3 ubiquitin ligase that translocates to altered mitochondria to increase the rate of phagosome formation. Parkin recruitment to mitochondria leads to ubiquitination of several targets including mitochondrial fusion proteins, such as mitofusins [85, 86], which promotes mitochondrial fragmentation prior to engulfment by the phagosome [87]. Evaluate the accumulation of mitophagic markers to alter mitochondria is an easy and well-established approach to study the activation of mitophagic response in the cells. With the subcellular fractionation protocol, here described, it is possible to obtain highly purified mitochondria fraction from cultured cells and liver, where analyze the expression levels of mitophagic markers accumulated. 3.2.1 Materials
1. Biological Material: cell lines of interest or liver biopsy. 2. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 3. FBS (ThermoFisher, 10270106). 4. Phosphatase inhibitor cocktail (100) (Merck, P0044). 5. Protease inhibitor cocktail (100) (Merck,P8340). 6. 1 l of Tris–HCl 1 M (pH 7.4): Dissolve 121.14 g of TrizmaBase in 500 ml of bidistilled water, adjust pH to 7.4 using HCl, bring the solution to 1 l with bidistilled water and store at 4 C. 7. ½ l of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) 0.5 M (pH 7.4): Dissolve 59.57 g of HEPES in 400 ml of bidistilled water, adjust pH to 7.4 using KOH, bring the solution to 500 ml with bidistilled water and store at 4 C. 8. 100 ml of 100 mM Ethylene-bis(oxyethylenenitrilo)tetraacetic acid (EGTA) (pH 7.4): Dissolve 3.8 g of EGTA in 70 ml of bidistilled water, adjust pH to 7.4 with KOH, bring the solution to 100 ml with bidistilled water and store at 4 C. 9. ½ l of Homogenization Buffer (HB) for Cells: (Composition: mannitol 225 mM, sucrose 75 mM, and 30 mM Tris–HCl, pH 7.4). Dissolve 20.5 g of mannitol, 13 g of sucrose in 400 ml of bidistilled water and add 15 ml of 1 M Tris–HCl (pH 7.4). Leave the buffer for about 30 min at 4 C to cool down. Check the pH of the buffer and adjust if necessary with KOH (if too low) or HCl (if too high) and bring the solution to a final volume of 500 ml with bidistilled water and store at 4 C. The buffer needs to be prepared fresh and must be free of protease and phosphatase inhibitor cocktails to avoid sample alteration. For liver homogenization: (composition: 225 mM
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mannitol, 75 mM sucrose, 0.5% BSA, 0.5 mM EGTA, and 30 mM Tris–HCl, pH 7.4). Dissolve in 150 ml of HB for cells 0.75 g of albumin and 0.75 ml of 100 mM EGTA (pH 7.4) and store at 4 C. 10. 100 ml of Mitochondria Isolation Buffer: (Composition: 250 mM mannitol, 5 mM HEPES (pH 7.4), and 0.5 mM EGTA). To prepare 100 ml of mitochondria isolation buffer, dissolve 4.56 g of mannitol in 80 ml of bidistilled water, add 1 ml of 0.5 M HEPES (pH 7.4) and 0.5 ml of 100 mM EGTA (pH 7.4). Check the pH of the buffer and adjust if necessary and bring the solution to a final volume of 100 ml with bidistilled water and store at 4 C. This buffer needs to be prepared fresh and must be free of protease and phosphatase inhibitor cocktails to avoid sample alteration (see Notes 1–3). 11. Cell culture dishes, 100 mm in diameter. 12. Cell scrapers (Sarstedt, 83.1830). 13. Stirrer motor with electronic speed controller (Cole-Palmer, EW-04369-10). 14. Motor-driven tightly fitting glass/Teflon Potter-Elvehjem homogenizer. 15. Loose- and tight-fitting glass Potter dounce homogenizer. 16. Oak Ridge Nalgene 30 ml tubes (for Sigma rotor angular 6 30 ml, Model 12139). 17. 1.5 ml Eppendorf microfuge test tubes (Eppendorf AG, 0030 120.086). 18. SW 40 rotor (swinging bucket, 6 14 ml, 40,000 rpm, 285,000 g) (Beckman, 331302). 19. Sigma rotor angular 6 30 ml (Merck, 12139). 20. Refrigerated Sigma low-speed centrifuge (Sigma (Braun), Model 2 K15, tabletop). 21. Low-speed MPW 342 centrifuge with rotor no. 12108. 3.2.2 Method
Solutions, centrifuge, rotor, centrifuge tubes, and homogenizer should be prechilled to 4 C. Carry out all procedures at 4 C or working in ice. For Adherent Cells: 1. Amplificate cells in petri dishes, to obtain at least two confluent petri for experimental conditions. In our experience, it is minimum number of plates useful to isolate a sufficient amount of mitochondria. 2. Discard the medium in culture dishes and wash the cells with ice-cold 1 PBS.
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3. Scrape the adherent cells using cell scraper, then collect the cells in centrifuge tube. 4. Centrifuge the cells at 400 g for 5 min at 4 C. Remove the supernatant and resuspend the pellet in 1 ml of ice-cold homogenization buffer. 5. Transfer the cells to glass/Teflon-dounce Potter homogenizer. 6. Homogenize the cells using a tight pestle of dounce potter, every 25 strokes, and control cell integrity under a light microscope (see Note 4). 7. Finish homogenization when 80–90% of cell damage has been attained. For Liver: 8. Rinse the murine liver twice with ice-cold 1 PBS. 9. Remove gallbladder and transfer the liver into a 50 ml tube. Wash the liver using ice-cold homogenization buffer for liver. 10. Discard the bloody homogenization buffer and repeat the wash for three times. 11. Using scissors, cut the liver in small pieces. 12. Collect the liver pieces and wash once again with ice-cold homogenization buffer for liver. 13. Discard the bloody homogenization buffer, transfer the liver pieces to the glass/Teflon Potter homogenizer. Add homogenization buffer for liver in the ratio 4 ml of buffer per gram of liver. 14. Homogenize the liver pieces using a pestle by eight strokes. Check the integrity of homogenized cells under a light microscope. Once Obtained the Homogenized Cells: 15. Collect an aliquot (homogenate fraction) for western blot analysis. Add to homogenate aliquot the protease and phosphatase inhibitor cocktails then store at 20 C. 16. Transfer the homogenized cells to 30 ml polypropylene centrifugation tubes, centrifuge at 600 g for 5 min at 4 C. 17. Transfer the supernatant to clean centrifuge tube and discard the pellet. 18. Repeat once again the centrifugation at 600 g for 5 min at 4 C. 19. Transfer the resulting supernatant to clean centrifuge tubes and discard the pellet. 20. Centrifuge the resulting supernatant at 15,000 g for 10 min at 4 C.
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21. Collect the supernatant (cytosolic fraction containing lysosomes and microsomes contamination) in a clean centrifuge tube to prepare the cytosolic fraction, while the pellet will be resuspended in mitochondria isolation buffer (see below). Preparation of the Cytosolic Fraction 22. Centrifuge the resulting supernatant at 17,000 g for 30 min at 4 C, to eliminate the lysosomes. 23. Collect the resulting supernatant in a clean tube, centrifuge at 100,000 g for 90 min at 4 C, to separate the cytosolic and ER fractions. 24. Collect the resulting supernatant (cytosolic fraction) and add the protease and phosphatase inhibitor cocktails for western blot analysis, then store at 20 C. 25. The pellet (ER fraction) may be resuspended in homogenization buffer per western blot analysis. Preparation of Mitochondrial Fraction 26. Gently wash the pellet with mitochondria isolation buffer without resuspending the pellet. 27. Gently discard the buffer, then resuspend the pellet by repeated pipetting in 1 ml of mitochondria isolation buffer. 28. Complete the resuspension of the mitochondrial pellet, using a loose pestle of dounce potter, ten strokes will be sufficient. 29. Centrifuge mitochondrial suspension at 15,000 g for 10 min at 4 C. 30. Discard the supernatant and resuspend gently the mitochondrial pellet in homogenization buffer for western blot analysis, adding the protease and phosphatase inhibitor cocktails, then store the mitochondria fraction at 20 C. 31. The quality of preparations will be checked by western blot, using different markers for a single fraction, to exclude the presence of contamination. Mitochondrial proteins and cytoskeleton proteins will be used as specific markers for mitochondrial and cytosolic fraction, respectively. For the protocol of western blot, please refer to Subheading 4.1.2. 3.2.3 Data Handling/ Processing
Excluded the presence of contaminations in the cytosolic and mitochondrial fractions, the intracellular redistribution to mitochondria of mitophagic markers, such as OPTN, NDP52, parkin, and PINK1, will be analyzed. In resting condition, these mitophagic markers are expressed in the cytoplasm, their redistribution and accumulation to mitochondrial fraction indicates the activation of mitophagic response (Fig. 5).
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-
+
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Mitochondria -
+
Parkin
GAPDH
pan/VDAC1
Fig. 5 Parkin translocation by subcellular fractionation. Parkin localization at the mitochondria was assessed by immunoblot after subcellular fractionation. Following treatment with the mitochondrial uncoupler FCCP, Parkin relocates from cytosolic to the mitochondrial fractions. GAPDH and pan/VDAC1 were used as cytosolic and mitochondrial markers, respectively 3.2.4 Notes
1. BSA is essential to bind and remove free fatty acids. 2. It is important to use sucrose containing low-Ca2+ contamination. Ca2+ contamination can provoke swelling of the mitochondria. 3. Prepare all buffers freshly on the day of the experiment to avoid sample alteration. 4. High force and speed during homogenization with pestle should be avoided to preserve mitochondria integrity.
3.3 Detecting and Quantifying Mitophagy in Human Body Fluids
Biomarkers can be assumed as multifaceted reporters of healthy status or pathological disorders. Typically, they help to evaluate prognosis or disease risk, to guide clinical diagnosis and to monitor individual response to treatment. Up to now, few studies have investigated the possibility of using mitophagy-related proteins as diagnostic and monitoring biomarkers for the most common human diseases. Recently, we demonstrated that both autophagic and mitophagic elements are present in human body fluids of patients affected by neurodegenerative disorders [88–90]. Most interestingly, in these studies, we demonstrated that the levels of mitophagic elements were related to the inflammatory status of the patient and the active/inactive state of the investigated disease. Considering this aspect, monitoring mitophagy by detecting its essential component in several body fluids of affected patients may be of critical importance. The ELISA is a well-established approach to test the presence of one or more antigens in a sample. Usually, for capturing the analyte to be measured, an antigen-specific antibody is precoated
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onto the bottom of the well. Then, a complex will form when the detection antibody (with biochemical modifications that can vary among manufacturers) and the enzyme substrate are added to the wells. This will produce a visible signal correlated to the amount of the analyte(s) present in the sample. ELISA assay is able to determine circulating levels of proteins involved in the mitophagic pathway such as Parkin, PINK1, OPTN, and NDP52 (CALCOCO2) from human body fluids and with great sensitivity (Parkin: 1 pg/ml and 0.1 ng/ml for PINK1, OPTN, and NDP52), high detection range (Parkin: 1–5000 pg/ ml; NDP52: 0.1–10 ng/ml; OPTN: 0.25–8 ng/ml, and for PINK1: 0.625–20 ng/ml), and excellence specificity with no significant cross-reactivity among protein analogs. 3.3.1 Materials
1. Human Parkinson disease 2/parkin (PARK2) ELISA kit, (MyBioSource, MBS732278 or similar). 2. Human serine/threonine-protein kinase PINK1, mitochondrial (PINK1) ELISA kit, (MyBioSource, MBS9327222 or similar). 3. Human optineurin (OPTN) ELISA kit, (MyBioSource, MBS069530 or similar). 4. Human calcium binding and coiled coil domain containing protein 2 (CALCOCO2) ELISA kit, (MyBioSource, MBS7220182 or similar). 5. Human Samples: serum and plasma, possible use of other biological fluids (saliva, urine, feces). 6. Refrigerated centrifuge. 7. Microplate reader (wavelength: 450 nm) with a dedicated software (i.e., SkanIt Software, ThermoFisher; SPECTROstar Nano, BMG labtech or compatible). 8. Incubator. 9. Precision single and multichannel pipette. 10. Disposable tips. 11. Clean tubes and Eppendorf tubes. 12. Deionized or distilled water. 13. All materials supplied by each kit.
3.3.2 Methods
Carry out all procedures at RT, unless otherwise specified. Consider the procedure listed below as the same for all ELISA kits proposed, unless otherwise specified. Sample Collection and Preparation (see Notes 1 and 2) 1. Serum: Place whole blood sample from 30 min to 2 h or put it at 4 C overnight and centrifuge for 15 min at approximately 1000 g.
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2. Plasma: Collect plasma using EDTA-Na2 as an anticoagulant. Centrifuge samples for 15 min at 1000 g at 4 C within 30 min of collection. 3. Other biological fluids (saliva, urine, feces). Centrifuge samples for 20 min at 1000 g at 4 C (see Notes 1 and 2). 4. Collect supernatant from 1–3 steps and estimate the concentration of target protein in the assay by making preexperiment readings. Then, select a proper dilution factor so that protein concentration falls in the optimal range of detection provided by the manufacturer. Dilution of the sample should be performed with the dilution buffer of the kit or PBS (pH 7.0–7.2) (see Notes 3 and 4). Reagents Preparation. Bring all components of the kit at RT before use. 5. Wash buffer. Dilute 10 ml of the provided wash buffer into 990 ml deionized or distilled water, mix gently. Solution can be stored at 4 C. 6. Other components are ready to use (included standards). Assay Procedure (see Notes 5 and 6) 1. Aliquot 0.1 ml of standards (in the case of PINK1 and OPTN, 0.05 ml) into the standard wells (see Note 7). 2. Aliquot 0.1 ml of PBS (pH 7.0–7.2) into the control well. 3. Aliquot 0.1 ml of diluted (or undiluted) sample (in the case of PINK1 and OPTN, 0.05 ml) into sample test wells (see Notes 2 and 7). 4. Aliquot 0.01 ml of balance solution into sample test wells only and mix well (if the sample is serum or plasma, this step could be skipped for both kits). 5. Add 0.05 ml of conjugate (in the case of PINK1 and OPTN, 0.1 ml) to each well (not to the control one) and mix well. 6. Cover the plate with a coverslip and incubate at 37 C for 60 min. 7. Remove the coverslip and discard standards, control, and samples. 8. Wash the plate five times with 0.3 ml wash buffer. 9. Add 0.05 ml substrate A plus 0.05 ml substrate B to each well (see Notes 1 and 8). 10. Seal the plate with a coverslip and incubate at 37 C for 15–20 min (see Note 9). 11. Add 0.05 ml stop solution into each well and mix them (see Note 1). 12. Read the optical density (O.D.) at 450 nm in a microplate reader immediately after the stop solution.
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3.3.3 Data Analysis
Set up the software of the microplate reader in order to: 1. Shake gently (5–10 s) the plate before acquisition. 2. Acquire absorbance for all experimental wells. 3. Calculate the average of duplicate readings for standards, control, and samples. 4. Subtract the average control O.D. from all other well readings before result interpretation. 5. Make a standard curve by plotting the average standard O.D. on the Y-axis against the concentration on the X-axis and draw a best fit curve using (if possible) a four parameter logistic (4-PL) method for Parkin and NDP52 detection and a linear curve fitting for PINK1 and OPTN measurements (Fig. 6) (see Note 10).
Fig. 6 Detecting and quantifying mitophagy in human body fluids Representative standard curve with a four parameter logistic (4-PL) method for Parkin detection (upper panel) and a linear curve fitting for PINK1 measurements (lower panel)
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6. Calculate samples concentration in pg/ml (Parkin) or in ng/ml (PINK1, OPTN, and NDP52) on the basis of the standard curve equation (see Figs. 1 and 2 as example) and multiply by the dilution factor (see Note 9). 3.3.4 Notes
1. Substances provided in the kit may be hazardous in case of skin and eye contact or inhalation. Be careful and wear gloves. 2. All blood components and biological materials should be handled as potentially hazardous. 3. The manufacturer and we suggest preexperiments with the following conditions: undiluted samples, diluted 1:2 and 1:4. 4. Samples should be frozen if not immediately analyzed; in that case, avoid also multiple freeze-thaw cycles. 5. Cover all kit components and store at 4 C when not in use. 6. Do not use reagents after the kit expiration date. 7. It is recommended that all conditions are run in duplicate. 8. TMB is light-sensitive, please avoid exposure to light. 9. At the end of the procedure if the color is not dark, you can prolong the incubation time until 30 min. 10. Coefficient of determination of the standard curve should be 0.98.
4
Measuring the Mitochondrial Balance Mitochondria are the main site for diverse biochemical processes, including the Krebs cycle [91], β-oxidation of fatty acids [92], oxidative phosphorylation [93], and Ca2+ homeostasis [94, 95]. To accomplish these physiological processes, mtDNA, lipids, and proteins occasionally become damaged and must be repaired. A large number of cellular pathways are involved to maintain their normal function and provide an efficient quality control system. During these events, mitochondria undergo several rearrangements, in particular in their morphology, functions, and metabolism [96]. To monitor, these dynamics represent a fundamental aspect of understanding the molecular mechanisms involved in the mitochondrial quality control system. Following, we will describe the main methods to assess parameters of mitochondrial function, metabolism, morphology, and protein damage.
4.1 Assessment of the Mitochondrial Structures
It is widely accepted that the fragmentation of the mitochondrial network represents the most common event following a stress condition. This parameter can be detected by measuring the length of a cell’s mitochondrial population. Fluorescent microscopy represents the main method to image the mitochondrial network and fragmentation events. This information can be obtained by
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transfecting cells with plasmids that present a mitochondrial target sequence. The most used are the mitochondrially targeted green fluorescent protein (mtGFP) plasmid [96] and the red fluorescent protein mCherry-Mito7 [80]. 4.1.1 Materials
1. Biological material (cell cultures of interest). 2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 3. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 4. Appropriate transfection reagent (e.g.; Lipofectamine LTX with Plus Reagent, ThermoFisher, 15338100). 5. mtGFP. 6. Glass coverslips 24 mm in diameter. 7. Six-well culture plates. 8. 4% PFA solution, pH 7.4, in 1 PBS. Add 4 g of PFA in 100 ml of 1 PBS. 9. Inverted Nikon confocal microscope system A1 R.
4.1.2 Methods
Sample Preparation and Transfection 1. Plate cells on glass coverslips at a density of 100,000 cells. 2. When cells reached a confluence of 50–60%, transfect them with 500 ng mtGFP plasmid by using the appropriate method (see Note 1). 3. After 4–6 h, replace the transfection medium with warm medium culture. 4. After 24 h, image the cells. Alternatively, cells can be fixed in 4% PFA for 15 min at RT, washed three times with 1 PBS, and stored at 4 C in the dark (see Note 2). Measurement: 5. Move the coverslips into a cell chamber and place in a 37 C thermo-controlled stage of the inverted confocal microscope system. 6. Capture picture at high-resolution by using standard binning mode 1 1 (meaning that each logical pixel is equal to one physical pixel) and collect a z-stack series of images at 0.2–0.5 μm each, to capture the whole mitochondrial network of the cell (see Note 3). 7. To obtain a statistical measurement of the mitochondrial network, acquire at least 20–25 images per condition and store images.
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Fig. 7 Assessment of the mitochondrial structures. Representative images showing the mitochondrial network derived from mtGFP-expressing cells. Left panel represents the volume rendering of a cell before denoising and subtraction of background (BG, middle panel). The right panel depicts the segmented image generated by the 3D objects counter plug-in installed in Fiji software 4.1.3 Data Handling/ Processing
4.1.4 Notes
The analysis of mitochondrial morphology may be achieved directly with the microscope imaging software or by using open-source software programs such as Fiji software (Fig. 7). We suggest using the plug-in 3D objects counter installed in the open-source Fiji software. An automated algorithm will calculate the average volume of mitochondria, the total volume, and the number of mitochondria counted for each cell. Before running the 3D objects counter plug-in, we recommend to denoise images by applying a Gaussian filter (Menu!Filters!Gaussian. . .) and remove the background setting the rolling ball to a size of 1 um (Menu Process ! Substract background. . .) (see Note 4). 1. Primary cell cultures are “hard to transfect” and alternative methods to transfection are needed. A large series of mitochondrial-specific fluorescent dyes (MitoTracker Green, MitoTracker Orange, MitoTracker Red, and MitoTracker Deep Red) may be used to stain the mitochondrial structures of primary cell samples. Please remember that these chemical fluorescent dyes are sensitive to alteration in mitochondrial membrane potential. 2. Several fixation methods are reported to alter mitochondrial structures. We recommend to fix cells with 4% PFA. 3. Attenuate laser excitation potency to minimize photobleaching and photodamage and keep constant camera settings and illumination across samples. 4. The background subtraction should not be modified between samples.
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4.2 Assessing Changes in Mitochondrial Functioning by Immunoblot of Mitochondrial Complexes
4.2.1 Materials
The method described in Subheading 4.1 is a useful approach to investigate the mitochondrial amounts by immunoblot. Despite this, it cannot provide information about the functionality of mitochondria. This can be done by measuring the levels of all five mitochondrial complexes of the electron transport chain (ETC). Notably, dysfunction in mitochondrial ETC compromises ATP production and accelerates the generation of free radicals. To evaluate the mitochondrial ETC function, the conventional methods used are spectrophotometric assays and blue native polyacrylamide gel electrophoresis (BN-PAGE) [97]. Alternatively, the protein complexes can be assessed by using commercially antibody cocktails that probe for all five complexes at once. 1. Biological material (cell cultures of interest). 2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 3. RIPA lysis buffer (150 mM sodium chloride, 1.0% NP-40 or Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 50 mM Tris, pH 8.0, supplemented of protease and phosphatase inhibitor). 4. CAPS-based transfer buffer (10 mM CAPS, pH 11, 10% methanol) (see Note 1). 5. Tris-buffered saline (TBS; 10): 1.5 M NaCl, 0.1 M Tris–HCl, pH 7.4. 6. TBS containing 0.05% Tween-20 (TBS-T). 7. Blocking Solution: 5% milk in TBS-T. 8. Precast gel 4–12% NW04120BOX).
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9. PVDF membranes. 10. Primary antibody anti-OXPHOS (see Notes 2 and 3). 11. Primary antibody anti-GAPDH. 12. Restore western blot stripping buffer (ThermoFisher, 21059). 4.2.2 Methods
Sample Preparation and Homogenates Collection 1. Plate cells in 6-well plates at a density of 150,000 cells. 2. After at least 24 h, cellular homogenates are collected and lysed by using RIPA lysis buffer. Keep homogenates in ice and vortex them every 5 min. 3. After 30 min, centrifuge cell lysates at 12,000 g for 10 min at 4 C, collect the supernatant, and keep them at 80 C. Western Blot for OXPHOS Components
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4. After having determined the protein concentration for the cell lysate, prepare 10–15 μg of each the sample in nondenaturating conditions. To avoid unspecific aggregation of the proteins, we recommend to do not boil the samples before. 5. Load samples in a precast gel 4–12% and perform the electrophoresis at 150 mV for 30–45 min. 6. Following electrophoresis, activate PVDF with methanol for 1 min and rinse with transfer buffer. Assemble the transfer stack with sponges, filter papers, PVDF membrane, and gel, being careful to that the gel is facing the cathode () and the membrane side is facing the anode (+). We recommend to transfer the gel for 2 h at 350 mA. 7. After the protein transfer, rinse the membrane in TBS-T and place the membrane in blocking solution and incubate with agitation for 1 h. 8. Place the blot in the primary antibodies solution (see Note 4) and incubate with agitation overnight. Wash the blot three times for 5 min. 9. Place the blot in the secondary antibody solution and incubate with agitation for 1 h at RT. Wash the blot three times for 5 min. 10. Proceed with chemiluminescent detection. Western Blot for Loading Marker 11. After detection, wash blot to remove chemiluminescent substrate. 12. Incubate blot in Restore western blot stripping buffer for 15 min at RT. 13. Wash the blot three times in TBS-T with agitation for 10 min at RT. 14. Place the membrane in blocking solution and incubate with agitation for 1 h. 15. Place the blot in the primary antibodies solution and incubate with agitation O.N. Wash the blot three times for 5 min. 16. Place the blot in the secondary antibody solution and incubate with agitation for 1 h at RT. Wash the blot three times in TBS-T with agitation for 10 min at RT. 17. Proceed with chemiluminescent detection (Fig. 8). 4.2.3 Data Handling/ Processing
Quantify the bands with a densitometric program (see Note 5) and then divide to the protein amounts of each complex with the loading marker. In the case two or more cellular conditions are analyzed, we suggest to normalize the values obtained with a mitochondrial marker, to calculate the quantities of the different proteins accordingly to the mitochondrial amounts.
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1. 10 mM CAPS Buffer pH: 11 will facilitate the protein migration in an electric field. Alternatively, use a lower pH buffer such as an acetic acid buffer. 2. This can be done individually by purchasing separate antibodies and probing each complex individually. 3. All mETC complexes are very sensitive to heating. Be careful to do not heat over 50 C. 4. The antibody cocktail (1.5 mg/mL) should be diluted 250 to a final working concentration of 6.0 μg/ml for western blotting. 5. COXI band will appear at ~35 kDa and not at its true molecular weight at 57 kDa.
4.3 Assessment of Mitochondrial Function Utilizing Seahorse Extracellular Flux
The metabolic activity of mitochondria may be detected by using the XF96 Extracellular Flux Analyzer (Seahorse Bioscience) [98– 101]. By adding specific modulators of respiratory complexes during the assay, this instrument permits to reveal key parameters of mitochondrial function: basal respiration (OCR-basal) that shows the energetic demand of the cell in baseline conditions; ATP-production (OCR-ATP) that shows the ATP produced by
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mitochondria; maximal respiratory capacity (OCR-MRC) that is indicative of the maximum rate of respiration that the cell can achieve; spare respiratory capacity (SRC-OCR) that represents how the cell responds to an energetic demand (Fig. 9). To achieve this information, the instrument performs subsequent injections of modulators of respiration into each well during the assay. The modulators are: the inhibitor of ATP synthase (Complex V) oligomycin, which decreases electron flow determining a decrease in OCR; the uncoupling agent FCCP that disrupts mitochondrial membrane potential and provokes an increase in OCR; a mixed solution composed of rotenone and antimycin-A (Rot-AA) that inhibits the Complex-III, resulting in a collapse of the mitochondrial respiration. In the following protocol, we will describe the method to measure the OCR in adherent cells by using the Agilent Seahorse XF Cell Mito Stress Test. 4.3.1 Materials
1. Biological material (cell cultures of interest) (see Note 1). 2. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 3. Seahorse XFe96 Analyzer (Agilent Technologies). 4. XFe96 MitoStress Test kit (Agilent Technologies, 103015100). The kit contains six pouches. Each pouch contains one each of oligomycin, FCCP, and rotenone/antimycin A (Rot-AA) (see Note 2). 5. Seahorse XF96 cell culture microplates (Agilent Technologies, 101085-004) (see Note 1). 6. Seahorse XF calibrant (Agilent Technologies, 100840-000).
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7. Seahorse XF assay medium modified DMEM (XF DMEM) (Agilent. 8. Technologies, 102365-100). 9. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 10. 1 mM sodium pyruvate. 11. 10 mM D-(+)-glucose. 12. 2 mM L-glutamine. 13. 0.1% crystal violet solution. 14. 4% PFA solution, pH 7.4, in 1 PBS. Add 4 g of PFA in 100 ml of 1 PBS. 15. Microplate spectrophotometer. 4.3.2 Method
The Day Before of the Assay: 1. Add 200 μl of Seahorse Bioscience calibrant, pH 7.4, to each well of a Seahorse Bioscience 96-well utility microplate (see Note 3). 2. Place sensor cartridge on top of the utility plate and store at 37 C without CO2 overnight and turn on instrument and start XF software to allow instrument to stabilize at 37 C. 3. Plate cells in a range of 5 104 cells per well in DMEM. We recommend to be sure that in the day of assay, cells are uniformly spread throughout the well. 4. Pipette 200 μl of the cell suspension into each well of 96-well microplate. At least two wells should lack cells to be used as a blank control. Put the seeded plate into a 37 C, 5% CO2 incubator to allow cells to grow overnight. The Day of Assay: 5. Warm XF DMEM media to 37 C, add 2 mM L-glutamine, 10 mM glucose, and 1 mM sodium pyruvate and adjust the pH of the media to 7.4. 6. Remove all but 25 μl of media from each well and add 175 μl of warm XF DMEM media, pH: 7.4. Place the plate in a 37 C incubator without CO2 for 1 h before running an assay. 7. Prepare the inhibitors in XF DMEM media at a concentration of 10x higher than the final dilution. Prepare 3 ml of 15 μM oligomycin, 3 ml of 10 μM FCCP, and 3 ml of 5 μM Rot-AA and add 25 μl of each solution into injection ports A, B, and C, respectively. Fill also A, B, and C injection ports of blank wells (see Note 4).
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8. Running the assay with the desired protocol. The XF96 analyzer required an input protocol composed of mix time, wait, and measure time. We suggest using a protocol composed of 3-min mix time, 3-min wait time and 3-min measure time for each point. 9. After the assay, the OCR data should be normalized to cell number or protein concentration. We suggest quantifying cell number by using crystal violet method. Fix cells with 4% PFA for 15 min at RT and stain cells with a 1% crystal violet solution for 15 min at RT. Measure the absorbance at 595 nm by using a microplate reader (see Note 5). 4.3.3 Data Handling/ Processing
OCR-Basal, OCR-ATP, OCR-MRC, and OCR-SRC can be calculated by using the following equations: 1. Basal Respiration: (Last rate measurement before first injection)—(nonmitochondrial respiration rate). 2. Maximal Respiration: (Maximum rate measurement after FCCP injection)—(nonmitochondrial respiration). 3. ATP Production: (Last rate measurement before oligomycin injection)—(minimum rate measurement after oligomycin injection). 4. Spare Respiratory Capacity: (Maximal respiration)—(basal respiration). Further parameters can be calculated: 5. H+ (Proton) Leak: (Minimum rate measurement after oligomycin injection)—(nonmitochondrial respiration). 6. Non-mitochondrial Oxygen Consumption: Minimum rate measurement after Rot-AA injection (Fig. 10). H+ Leak represents the remaining basal respiration not coupled to ATP production and can be related to mitochondrial damage. Non-mitochondrial respiration may indicate the existence of subcellular enzymes that consume oxygen after the third injection of Rot-AA.
4.3.4 Notes
1. Bioenergetic information can be also assessed in isolated mitochondria and in nonadherent cells. 2. The protocol outlined here is for the 96-well format of the instrument. Volumes will need to be adjusted if another format is used. 3. Verify that the calibrant solution level is high enough to keep the sensors submerged. 4. Each series of ports (e.g., all ports A) must contain the same volume.
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Fig. 10 Oxygen consumption rate (OCR) in Mito Stress assay. Oxygen consumption rate (OCR) assessed in primary cells exposed to hypoxic condition. To perform these experiments, before measurements cells were incubated with a solution 50 μM cobalt (III) chloride hexahydrate (CoCl2). OCR was measured under basal condition and following perfusion of oligomycin, FCCP, and Rot-AA. Each data point represents an OCR measurement performed in at least three different wells of a 96-well plate. The graphs are representative of the calculated parameters for basal respiration (OCR-basal), ATP production (ATP-OCR), maximal respiration (MRC-OCR), spare respiratory capacity (SRC-OCR), proton leak (H + Leak), and non-mitochondrial respiration (non-mito OCR)
5. If it is not convenient to proceed with the protein concentration assay, it is possible to freeze the whole plate at 20 C until analysis. 4.4 Monitor Mitochondrial Turnover with MitoTimer
First efforts to measure mitochondrial turnover were performed in late 1950s by monitoring 35S-methionine incorporation into newly synthesized proteins [102]. Newly, a protocol describes the possibility to measure the changes of mitochondrial proteins by using deuterium labeling and mass spectrometry [103]. The main
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limitation of these methods is the impossibility to imaging the process. To visualize mitochondrial turnover, the fluorescent Timer construct (DsRed1-E5) has been recently targeted to the mitochondrial matrix by fusing the mitochondrial targeting sequence of the COX8A subunit [104]. The main characteristic of Timer is to shift its fluorescence overtime from green to red as the protein matures. In view of this, different information can be achieved by using this novel construct: 1. Increases in red signal without changes in green channel may suggest impairments in mitochondrial degradation. 2. The newly synthesized mitochondria appear green-only. 3. Yellow mitochondria are indicative of an intermediate stage of mitochondria and represent the fusion process of newly synthesized (green) and old/mature (red) mitochondria. 4.4.1 Materials
1. Biological material (cell cultures of interest). 2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 3. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 4. 4% PFA solution, pH 7.4, in 1 PBS. Add 4 g of PFA in 100 ml of 1 PBS. 5. Transfection reagent (Lipofectamine LTX with Plus Reagent, ThermoFisher, 15338100). 6. Mitochondrial fluorescent Timer construct (MitoTimer). 7. Glass coverslips. 8. Six-well culture plates. 9. Cell chamber (Attofluor cell chamber, ThermoFisher, A7816). 10. Inverted Zeiss LSM 510 confocal microscope (see Note 1).
4.4.2 Methods
Sample Preparation and Transfection 1. Plate cells onto coverslips (24 mm in diameter) at a density of 100,000 cells per coverslip. 2. When cells reached a confluence of 50–60%, transfect them with 1 μg/coverslips of MitoTimer plasmid by using the appropriate method. 3. After 4–6 h, replace the transfection medium with warm medium culture, and after 24 h cells image the cells. Alternatively, cells can be fixed in 4% PFA for 15 min at RT, washed three times with 1 PBS, and stored at 4 C in the dark (see Note 2).
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4. Move the coverslips into a cell chamber and place in a 37 C thermo-controlled stage of the inverted Zeiss LSM 510 confocal microscope (see Note 3). Fluorescence images were captured providing excitation at 488 nm (green) and 546 nm (red) with detection of green (500–540 nm) and red (580–640 nm) fluorescence signals (see Note 4). 5. To obtain a statistical measurement of the mitochondrial network, acquire at least 20–25 images per condition (see Note 5). 4.4.3 Data Handling
4.4.4 Notes
Image processing may be achieved by using the microscope software (ZEN elements for Zeiss instruments) or Fiji software. After background subtraction on both green and red channel, ratiometric images were generated, and the quantification of red/green ratio signal was performed by registering the mean pixel intensity. To quantify the yellow signal, we suggest to use the colocalization plug-in available in Fiji software or the colocalization command of ZEN software (Fig. 11). 1. MitoTimer is also suitable for analysis by flow cytometry using a 488 nm laser for excitation and detection in the FITC and PE channels. 2. Sample transfected with MitoTimer can be fixed. In addition, fixation prevents postfixation maturation and stabilizes green and red conformations. 3. Changes in temperature may provoke transition from green to red fluorescence. It is recommended to use a 37 C thermocontrolled stage during imaging. 4. Prolonged light exposure during live-cell imaging may accelerate the maturation (red photoconversion). Keep laser potency and exposure time low. 5. Keep exposure time and illumination constant across samples.
4.5 Monitor Mitochondrial Energetic Levels with Fluorescent Probes
Among the different major events that occur in mitochondria during damage/stress condition, the loss of mitochondrial transmembrane potential (ΔΨm) is the most significant [105]. This force is generated by proton pumps (C-I, III, and IV of mitochondrial ETC), and together with the proton gradient (ΔpH) is the central mitochondrial parameter that regulates ATP, synthesis, respiratory rate, and production of ROS. In addition to this, ΔΨm represents the driving force that regulates the transport of different charged compounds (such as Ca2+) [106], and its dissipation determines the activation of the mitophagic pathway regulated by PINK1-Parkin axis. Several fluorescent probes are used to investigate the levels of ΔΨm. Among them, the most used are fluorescent cationic dyes including rhodamine 123 (Rh123), JC1, tetramethylrhodamine ethyl (TMRE) or methyl (TMRM) ester, nonyl acridine orange
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Fig. 11 Monitor mitochondrial turnover with MitoTimer. Fluorescent images of MitoTimer expressing cells. Red-only mitochondria are representative of old mitochondria. Green-only mitochondria represent newly synthesized mitochondria. Yellow staining depicts an intermediate stage of mitochondria
(NAO), and merocyanine 540 [107]. From first glance, JC1 may represent the best opportunity due to the fact that this probe forms red-fluorescent aggregates in low-potential mitochondria or greenfluorescent aggregates in mitochondria characterized by highpotential. At the same time, this probe has diverse adverse effects like photodamage and changes of fluorescence independent from ΔΨm. In light of this, we suggest using TMRM or TMRE to monitor variation in ΔΨm. This cell-permeable fluorescent indicator accumulates in the mitochondrial matrix in proportion to ΔΨm, and compared to other probes, this has limited effect on mitochondrial respiration as well as low photobleaching and phototoxicity [108, 109].
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4.5.1 Materials
1. Biological material (cell cultures of interest). 2. Culture medium Dulbecco’s Modified Eagle Medium (DMEM; e.g., Gibco 11965084) supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. 3. 1 PBS buffer, pH 7.4, no phenol red. In 1 l of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, 0.24 g of KH2PO4. Adjust the pH to 7.4 with HCl. 4. Tetramethylrhodamine, TMRM (ThermoFisher, T668). 5. 10 mM FCCP. 6. Glass coverslips (24 mm in diameter). 7. Six-well culture plates. 8. Cell chamber (Attofluor cell chamber, ThermoFisher, A7816). 9. Inverted Zeiss LSM 510 confocal microscope.
4.5.2 Methods
Sample Preparation 1. Plate cells on glass coverslips (24 mm in diameter) a density of 150,000 cells. After seeding the cells, wait for at least 24 h (see Note 1). 2. Wash cells with prewarmed 1 PBS and incubate cells with TMRM in the appropriate cellular media at 37 C for 30 min. We recommend to use low concentration of the probe (10–25 nM range) to avoid autoquenching (see Note 2). 3. Move the coverslips into a cell chamber. 4. Place the coverslip on a stage incubator of the confocal microscope equipped with a high magnification objective (see Notes 3 and 4). 5. Once having adjusted the focus of the cells using reflected light and examine TMRM fluorescence by excitation at 515 nm and emission at 595 nm, collect images with time-lapse interval of 50 or 200 ms image acquisition/illumination time for 1 min to register the basal TMRM fluorescent intensity. 6. Apply a stimuli 1 μM FCCP, which will depolarize the ΔΨm. These changes provoke a decrease in TMRM fluorescence intensity and will be indicative of background fluorescent intensity of TMRM. Record images for 1 min (see Note 5).
4.5.3 Data Handling Analysis
Data analysis should be achieved by using the ROI tool from the microscope program or from Fiji. Select ROIs from mitochondrial regions and calculate the average fluorescence intensities. Subtract the average fluorescent intensity after FCCP addition from average basal fluorescent intensity of TMRM. Then, use Prism or Excel program to generate the graphs and plots indicative of changes in fluorescence intensity overtime (Fig. 12).
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Fig. 12 Monitor mitochondrial energetic levels with fluorescent probes. Representative trace of TMRM fluorescence signal showing calculable parameters. After registering the basal fluorescence intensity, FCCP was added to collapse the mitochondrial membrane potential (ΔΨm) and obtain the background fluorescent intensity. Images of fluorescence confocal microscopy of cells loaded with TMRM before and after FCCP addition
4.5.4 Notes
1. At low concentrations of mitochondria, the greater fraction of TMRM remains in the media. In this case, the method may be less sensitive to detect changes in ΔΨm. A mitochondrial protein concentration >1 mg/ml is predicted to be optimal for TMRM assay [108]. 2. TMRM staining must not be washed out. 3. Changes in temperature may provoke transition from green to red fluorescence. It is recommend to use a 37 C thermocontrolled stage during imaging. 4. We suggest to use a 40x magnification to capture an optimal number of cells and to keep laser excitation at low potency to minimize photobleaching and photodamage. 5. Keep exposure time and illumination constant across samples.
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Acknowledgments PP is grateful to Camilla degli Scrovegni for her continuous support. All authors thank the Associazione Ricerca Oncologica Sperimentale Estense (A-ROSE). Funding: PP was supported by Telethon (GGP15219/B), the Italian Association for Cancer Research (AIRC: IG- 23670), and by local funds from the University of Ferrara. CG was supported by local funds from the University of Ferrara, the Italian Association for Cancer Research (AIRC: IG19803), the Italian Ministry of Health (GR-2013-02356747), and European Research Council Grant 853057-InflaPML. SP was supported by Fondazione Umberto Veronesi. References 1. Patergnani S, Missiroli S, Marchi S, Giorgi C (2015) Mitochondria-associated endoplasmic reticulum membranes microenvironment: targeting Autophagic and apoptotic pathways in cancer therapy. Front Oncol 5:173 2. Giorgi C, Missiroli S, Patergnani S, Duszynski J, Wieckowski MR, Pinton P (2015) Mitochondria-associated membranes: composition, molecular mechanisms, and physiopathological implications. Antioxid Redox Signal 22:995–1019 3. Giorgi C, Danese A, Missiroli S, Patergnani S, Pinton P (2018) Calcium dynamics as a machine for decoding signals. Trends Cell Biol 28:258–273 4. Pinton P, Leo S, Wieckowski MR, Di Benedetto G, Rizzuto R (2004) Long-term modulation of mitochondrial Ca2+ signals by protein kinase C isozymes. J Cell Biol 165:223–232 5. Shimizu S (2019) Organelle zones in mitochondria. J Biochem 165:101–107 6. Saha PP, Vishwanathan V, Bankapalli K, D’Silva P (2018) Iron-sulfur protein assembly in human cells. Rev Physiol Biochem Pharmacol 174:25–65 7. Jiang S, Park DW, Stigler WS, Creighton J, Ravi S, Darley-Usmar V, Zmijewski JW (2013) Mitochondria and AMP-activated protein kinase-dependent mechanism of efferocytosis. J Biol Chem 288:26013–26026 8. Gomes LC, Di Benedetto G, Scorrano L (2011) During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat Cell Biol 13:589–598 9. Liu L, Feng D, Chen G, Chen M, Zheng Q, Song P, Ma Q, Zhu C, Wang R, Qi W et al
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Chapter 10 Analysis of Proapoptotic Protein Trafficking to and from Mitochondria Ignacio Vega-Naredo, Gabriela Oliveira, Teresa Cunha-Oliveira, Teresa Serafim, Vilma A. Sarda˜o, and Paulo J. Oliveira Abstract Mitochondria play a key role in cell death and its regulation. The permeabilization of the outer mitochondrial membrane, which is mainly controlled by proteins of the BCL-2 family, is a key event that can be directly induced by different signaling pathways, including p53-mediated, and results in the release of proapoptotic factors to the cytosol, such as cytochrome c, second mitochondria-derived activator of caspases/direct inhibitor-of-apoptosis (IAP) binding protein with low pI (SMAC/Diablo), Omi serine protease (Omi/HtrA2), apoptosis-inducing factor (AIF), or endonuclease G (Endo-G). Hence, the determination of subcellular localization of these proteins is extremely important to predict cell fate and elucidate the specific mechanism of apoptosis. Here we describe experimental protocols that can be used to study the subcellular location of different proapoptotic proteins to be used in basic cell biology and toxicology studies. Key words Mitochondria, Proapoptotic proteins, Cell fractions, Immunoblot, Immunoprecipitation, Immunocytochemistry
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Introduction Mitochondria are cellular organelles with important functions in cell life and death, acting as cellular powerhouses by producing energy to maintain cellular activity. However, mitochondria are also important checkpoints for cell fate decisions, playing a crucial role in programmed cell death (PCD) pathways. The permeabilization of the outer mitochondrial membrane (OMM) is a fundamental step in several tightly regulated pathways of cell death, allowing the release of proteins that are usually only present in the intermembrane space, and signaling cell death programs [1]. One of the best-characterized types of cell death is apoptosis. Apoptotic signals may originate outside the cell (extrinsic pathway) or from any
Carlos M. Palmeira and Anabela P. Rolo (eds.), Mitochondrial Regulation: Methods and Protocols, Methods in Molecular Biology, vol. 2310, https://doi.org/10.1007/978-1-0716-1433-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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intracellular compartment (intrinsic pathway), constituting two distinct yet complementary apoptotic mechanisms. Both intrinsic and extrinsic stimuli may lead to OMM permeabilization. The OMM is selectively permeable to solutes, and its permeability and integrity are regulated by proteins of the BCL-2 family [2]. This family includes pro- and antiapoptotic members, and depending on their function and the presence of different BCL2 homology (BH1-4) can be divided into three groups [3, 4]: antiapoptotic proteins, such as BCL-2 and BCL-xL, that have four different BH domains—BH1-4 and pro-apoptotic pore-forming proteins, such as BAX and BAK, that are also multidomain proteins with three different BH domains—BH1-3. BH3-only proteins, that have proapoptotic activity, have only one BH domain and can exert their proapoptotic function either by facilitating or by activating BH1-3 proteins, which then initiate OMM permeabilization. Facilitators or de-repressors, such as BAD, interact with BH1–4 proteins, dissociating them from sequestered proapoptotic proteins, which become free to promote OMM permeabilization. The activators, such as tBID (which results from the cleavage of BID by caspase-8), directly activate BH1-3 proteins, either by stimulating the translocation of Bad to the OMM or by interacting with BAK. OMM permeabilization may occur through a Bax/Bakmediated mechanism [5] or by the opening of the mitochondrial permeability transition pore (MPTP) in the inner mitochondrial membrane [6]. In the latter mechanism, the opening of the MPTP can lead to the mechanical rupture of the OMM, followed by a release of proapoptotic proteins or, instead, to the recruitment of proapoptotic proteins to the OMM, causing mitochondrial depolarization [7]. p53 is a redox-sensitive transcription factor with a broad range of actions, some of them related to survival and cell death [8, 9]. In general, the tumor suppressor p53 exerts important roles in cell cycle progression and cell death by coordinating multiple options for cellular response to genotoxic stress. p53 inhibits replication of the genome by blocking cell cycle progression at a G1/S checkpoint in response to DNA damage [10]. In unstressed cells, p53 protein levels are kept low, but increase following stress signals acting through both transcription-dependent and -independent mechanisms to coordinate the appropriated cellular responses [11]. p53 activity depends on its ability to activate or repress gene transcription. Thus, p53 oscillates between latent and active sequence-specific DNA binding conformations and is differentially activated through posttranslational modifications including phosphorylation, acetylation, ubiquitination, sumoylation, among others [12, 13]. On the other side, non-sequence-specific DNA binding may mediate other p53 actions [14]. In addition, p53 is also involved in mitochondrial-dependent cell death, collaborating
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in the execution of the apoptotic pathway. In this context, p53 undergoes a nuclear–cytoplasm–mitochondria trafficking and Western blotting and immunocytochemistry are usually performed to detect the presence of p53 in nuclear, cytoplasmic, and mitochondrial extracts isolated as described in this chapter. The analysis of immunoreactive bands in nuclear fractions indicates whether nuclear translocation is occurring. The nuclear localization is critical for its transcriptional activity by activating genes that arrest cell growth and repair DNA damage. To further confirm its transactivation function, the evaluation of the expression of the transcription target genes of p53 such as PUMA, NOXA, BAX, BID, and DRAM is often useful [15]. As described above, the activity of the p53 gene product is regulated by posttranslational modifications. These modifications of p53 affect its stability and can be a potential mechanism to select the target genes conferring differential binding affinity to the response elements. For example, acetylation can directly affect p53 activity by enhancing its transcriptional activity [16], which can be detected using different acetyl-p53 antibodies available. Acetylation of p53 at carboxyl-terminal lysine residues enhances its transcriptional activity associated with cell cycle arrest and apoptosis. However, p53 acetylation at Lys-320/Lys-373/ Lys-382 is also required for transcription-independent functions involving BAX activation [17]. Furthermore, structural studies showed that acetylation at Lys-120 promotes a switch on the p53 protein conformation, increasing its ability to bind to the BAX binding site [18]. The phosphorylation of p53 at Ser15 and Ser20 following DNA damage can also be detected by immunoblotting to evaluate its transactivation function since this phosphorylation promotes p53 activation and stabilization reducing the interaction between p53 and its negative regulator, the oncoprotein murine double minute 2 (MDM2) [19]. In addition, phosphorylation of p53 in Ser392 influences its transcriptional activation, regulates its oncogenic function, and is also involved in the regulation of p53 mitochondrial translocation and transcription-independent apoptosis [20–22]. MDM2 inhibits cytoplasmic retention of p53 by targeting it for ubiquitination and proteasomal degradation [23]. Some reports suggest not only that the cytoplasmic retention of p53 can repress autophagy [24, 25] but also that this retention can be mediated by acetylation, since p53 acetylation reduces its ubiquitination status [26]. Although the precise molecular mechanisms behind this remain unclear, p53 deacetylases may be upregulated to mediate ubiquitination and degradation of p53 [27]. Because of this, the immunoblot analysis of cytoplasmic extracts can reveal whether p53 is accumulated in cytoplasm and can help to reveal the function of p53 in a specific context. p53 can promote cell death independently of transcription by two different mechanisms, each of which is
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assigned to a specific localization of the protein: cytosol or mitochondrial. Both modes of action converge in the permeabilization of the OMM via activation of the proapoptotic proteins BAX or BAK. In fact, cytosolic p53 can directly activate BAX and thereby induce apoptosis [28]. On the other hand, in response to a broad spectrum of apoptotic stimuli, a pool of p53 translocates to mitochondria and triggers a direct mitochondrial p53 death program [29]. For this, p53 physically interacts with the BCL-2 family member proteins BCL-xL and BCL-2 antagonizing their antiapoptotic function and inducing OMM permeabilization [30]. Furthermore, mitochondrial p53 directly promotes the proapoptotic activities of BAX and directly induces BAK oligomerization [31]. p53 also interacts with the antioxidant enzyme superoxide dismutase 2 (SOD2) leading to a reduction of its superoxide scavenging activity, and a subsequent decrease of mitochondrial membrane potential which contributes to the induction of proapoptotic mitochondrial alterations [32]. Surprisingly, p53 is released from mitochondria mediating a retrograde signaling pathway to the nucleus [33]. Therefore, the study of the subcellular localization of p53, its posttranslational modifications, and the levels of p53-related/-targeted proteins are crucial to examine the p53 pathway and elucidate its particular role. As described above, OMM permeabilization resulting from stress stimuli can result in the simultaneous release of proapoptotic factors that are normally limited to the mitochondrial intermembrane or intercristae space, including cytochrome c, apoptosisinducing factor (AIF), endonuclease G (endoG), Smac/Diablo, and Omi/HtrA2 [34]. Cytochrome c was first identified as being involved in mitochondrial bioenergetics, essential for the ATP production by oxidative phosphorylation, and later to apoptosis. Following the release of cytochrome c from the intermembrane space, it binds to apoptotic protease activating factor 1 (APAF-1) and dATP [35], forming the apoptosome complex and initiating cell death with the activation of caspase-9 and consequent triggering of the caspase cascade [36]. The second mitochondrial-derived activator of caspase (Smac), also known as direct IAP-binding protein with a low pI (Diablo) [36], resides in mitochondria in a mature form and is released during apoptosis, which promotes caspase-dependent apoptosis by controlling the activity of the inhibitor of apoptosis protein (IAP). IAPs are cytosolic proteins that inhibit caspase-3 and -7, and thus inhibit apoptosis [37]. Omi is a serine protease that, similarly to Smac, is also involved in the neutralization of IAPs. It can be released from mitochondrial intermembrane space into the cytoplasm upon apoptotic insult and cleaves both IAPs and cytoskeletal proteins, contributing to apoptosis in caspase-dependent and -independent manner [38].
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Fig. 1 Intrinsic Apoptotic Pathway. The intrinsic or mitochondrial apoptotic pathway initiates with mitochondrial membrane permeabilization, which may result from p53 signaling, with the formation of different pores in the mitochondrial membrane, such as Bax/Bak or mitochondrial permeability transition pore (MPTP), in the latter case by leading to OMM rupture or by triggering proapoptotic protein translocation to mitochondria following mitochondrial depolarization. Consequently, the release of apoptotic factors to the cytosol, including cytochrome c, second mitochondria-derived activator of caspases/direct inhibitor-of-apoptosis binding protein with low pI (SMAC/Diablo), Omi serine protease (Omi/HrtA2), endonuclease G (endoG), and apoptosis-inducing factor (AIF), occurs. The released cytochrome c reacts with deoxy ATP (dATP) and apoptotic protease activating factor 1 (APAF-1), leading to apoptosome formation. The apoptosome recruits procaspase-9, activating the initiator caspase-9, and the caspase cascade begins
Independently of caspase activation, cell death can occur with the release of endoG and AIF. EndoG is also localized in the mitochondria, and there is evidence that it resides within the intermembrane space and also bound to the inner membrane [39]. Due to mitochondrial membrane potential loss, the mature endoG is translocated to the nucleus and initiates oligonucleosomal DNA fragmentation. EndoG can regulate several mitochondrial enzymes expression, such as complex I (ND1 and ND2), complex IV (COX2), and complex V (ATPase6) [38] (Fig. 1). The AIF is a flavoprotein harboring NADH oxidase activity. Initially, the AIF was identified as a proapoptotic protein, inducing a type of programmed cell death independently of caspases activation. When AIF is added to purified nuclei extracts in a cell-free system, chromatin condensation and large-scale DNA fragmentation to 50 kbp fragments occur [40]. Those effects were observed in intact cells after diverse apoptotic stimuli and also in models of retinal
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degeneration, brain damage induced by hypoglycemia or ischemia, or myocardial infarction [41]. The AIF protein is encoded by a nuclear gene, synthesized in the cytoplasm and transported to mitochondria by the general import pathway, due to the mitochondrial localization sequence (MLS) in the precursor protein. Once in the mitochondrial intermembrane space, the MLS is proteolytically cleaved, the protein refolds and incorporates flavin adenine dinucleotide (FAD) cofactor, generating the mature protein. The oxidoreductase activity, dependent on the presence of the prosthetic group, is not critical for the apoptogenic effect of AIF but the structural parts of the oxidoreductase domain are necessary to the DNA binding [40]. A decrease in AIF enzymatic activity or a decrease in AIF expression results in decreased oxidative phosphorylation and increased free radical generation [42]. On the other hand and depending on cell type and cell death stimuli, AIF protein is involved in programmed cell death. The basic mechanism of AIF-induced cell death consists of AIF release from the mitochondrial intermembrane space to the cytosol and then on its translocation to the nucleus. Once in the nucleus, AIF binds to DNA and induces chromatin condensation and large-scale DNA fragmentation. The nuclear apoptosis induced by AIF requires direct interaction of AIF with DNA [43]. In the cytosol, AIF also promotes a decrease in mitochondrial ΔΨ, release of cytochrome c, and further AIF release from mitochondria, promoting a positive feedback amplification loop [40, 41]. In order to be released from the mitochondrial intermembrane space, AIF must be cleaved in a specific region to release the protein binding to the mitochondrial inner membrane. This cleavage is performed by proteases such as calpains and cathepsins that may have access to the mitochondrial intermembrane space during apoptotic stimuli and require the presence of calcium. The truncated AIF (tAIF) is then free to execute its caspase-independent apoptotic action. An interesting aspect is the fact that binding of AIF to DNA induces large-scale DNA fragmentation, but AIF itself does not possess DNAse activity. Thus, it has been proposed that the DNA-degrading capacity of AIF could be due to the recruitment of downstream nucleases, such as cyclophilin A (CypA) [44, 45]. In this chapter, we describe the material and the methods followed by us and other authors to study the proapoptotic protein trafficking to and from mitochondria; immunoblotting performed with different cellular fractions and immunocytochemistry using the antibodies are described in Table 1. Immunocytochemical approaches can be used to study the subcellular localization of proapoptotic factors described in this chapter. To evaluate their mitochondrial localization, co-localization assays using antibodies constitute an effective alternative (Fig. 2). One option is to develop
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Table 1 List of antibodies for western blotting (WB) and immunocytochemistry (ICC)
Antibodies
MW (kDa)
Company (Cat. No.)
Species crossreactivity
p53
53
Cell Signaling (2524)
H, M, R, Mk
Mouse IgG1
WB (1:1000); ICC (1:250)
p53
53
Santa Cruz (sc-6243)
H, M, R
Rabbit IgG
WB (1:500); ICC (1:50)
Phospho-p53 (Ser15)
53
BioVision (3515) H, M, R
Rabbit IgG
WB (4 μg/ml)
Phospho-p53 (Ser15)
53
Cell Signaling (9298)
H, M, R, Mk
Rabbit IgG
WB (1:1000); ICC (1:250)
Phospho-p53 (Ser329)
53
Cell Signaling (9281)
H, M
Rabbit IgG
WB (1:1000)
Acetyl-p53 (Lys379)
53
Cell Signaling (2570)
H, M
Rabbit IgG
WB (1:1000)
Puma
23
Cell Signaling (4976)
H
Rabbit IgG
WB (1:1000)
Puma
23
Cell Signaling (7467)
M, R
Rabbit IgG
WB (1:1000)
Noxa
15
Santa Cruz (sc-56,169)
H, M
Mouse IgG1
WB (1:500)
Bax
20
Cell Signaling (2772)
H, M, R, Mk
Rabbit IgG
WB (1:1000)
Bak
25
Cell Signaling (3814)
H, M, R, Mk
Rabbit IgG
WB (1:1000)
Bid
22 Cell Signaling (15) (2002)
H
Rabbit IgG
WB (1:1000)
Bid
22
Cell Signaling (2003)
M
Rabbit IgG
WB (1:1000)
DRAM
33
Rockland (600–401A70)
H, M, R
Rabbit IgG
WB (4 μg/ml); ICC
MDM2
90 Santa Cruz (60) (sc-965)
H, M, R
Mouse IgG1
WB (1:500); ICC (1:50)
SOD2
25
Santa Cruz (sc-18,504)
H, M, R
Goat IgG
WB (1:500); ICC (1:50)
Cell Signaling (3933)
H, M, R, Mk
Rabbit IgG
WB (1:1000)
Cell Signaling (2870)
H, M, R, Mk
Rabbit IgG
WB (1:1000)
Ubiquitin Bcl-2
26
Isotype
Applications (Recommended dilution)
(continued)
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Table 1 (continued)
Antibodies
MW (kDa)
Company (Cat. No.)
Species crossreactivity
Bcl-xL
30
Cell Signaling (2764)
H, M, R, Mk
Rabbit IgG
WB (1:1000); ICC (1:200)
Tom20
20
Santa Cruz (sc-11,415)
H, M, R
Rabbit IgG
WB (1:500); ICC (1:100)
Cytochrome c
12
Abcam (ab13575) M, R, H, P, Hr, Ff
Mouse IgG
WB (1:500); ICC (1:200)
Cytochrome c
12
Abcam (ab110325
M, R, C, H, Ce
Mouse IgG
WB (0.5 μg/ml); ICC (1 μg/ml)
Smac/Diablo
21
Cell signaling (2954)
H, Mk
Mouse IgG
WB (1:1000); ICC (1:100)
Apaf-1
130
Millipore (AB16503)
H, R, M
Rabbit IgG
WB (1:1000)
Omi
48
Abcam (ab33041) M, H
Mouse IgG
ELISA; WB (1:1000)
AIF
57
Santa Cruz (sc-13,116)
Mouse gG2b
WB (1:1000); ICC (1:100)
H, R, M
Isotype
Applications (Recommended dilution)
Anti-rabbit IgG-AP
Santa Cruz (sc-2007)
Goat IgG
WB (1:5000)
Anti-goat IgG-AP
Santa Cruz (sc-2022)
Donkey IgG
WB (1:5000)
Anti-mouse IgG1-AP
Santa Cruz (sc-2066)
Goat IgG
WB (1:5000)
Anti-rabbit IgG-FITC
Santa Cruz (sc-2012)
Goat IgG
ICC (1:400)
Anti-mouse IgG-TR
Santa Cruz (sc-2781)
Goat IgG
ICC (1:400)
Ce Caenorhabditis elegans, C cow, Ff fruit fly, Hr horse, H human, Mk monkey, M mouse, P pigeon, R rat
immunocytochemistry using a secondary antibody conjugated with a fluorochrome such as FITC and to label mitochondria using mitochondria-selective probes such as MitoTracker Red. Another option consists in a double immunofluorescence assay using a mixture of two primary antibodies, for example, one against the mitochondrial marker TOM20 and the other against the desired proapoptotic protein, using their respective secondary antibodies which have to be raised in different species and conjugated with two different fluorochromes (e.g., FITC-conjugated against rabbit and Texas Red-conjugated against mouse).
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Fig. 2 Representative micrographs of fluorescence microscopy of immunocytochemistry-labeled apoptotic proteins. Images show H9c2 cardiomyoblasts treated with the anticancer agent doxorubicin (DOX) and imaged for p53 (A–C) and Bax (D–G). Panels A–C show H9c2 cells treated with vehicle (A), 0.5 μM (B), and 1 μM (C) DOX for 24 h. After treatment, H9c2 cells were fixed with ice-cold methanol (which removed nuclear bound DOX), labeled with an antibody against p53. Epifluorescence microscopy images in panels A–C illustrate increased p53 nuclear labeling (in blue) after H9c2 cells treatment with DOX. Cells were observed by epifluorescence microscopy using a Nikon Eclipse TE2000U microscope equipped with a 40 Plan Fluor 1.3 NA oil immersion DIC objective and images were processed using MetaMorph software (Universal Imaging, Downingtown, PA). Panels D-G show confocal microscopy images of H9c2 cells labeled with Hoechst 33342 (nucleus, blue and pink), MitoTracker Red (mitochondria, red), and Bax (green). Confocal microscopy of H9c2 cells treated with 0 (D), 0.5 μM (E, G), and 1 μM (F) DOX for 24 h. After treatment, cells were fixed in paraformaldehyde (to maintain the integrity of the mitochondrial network) and subsequently immunolabeled with an antibody against Bax. Increased immunolabeling for Bax is observed after DOX treatment, especially forming large clusters in the cytosol of treated cells. This is clearly visible in panel G, obtained with a higher magnification. Images in panels D–G were obtained by using a Nikon C-1 laser scanning confocal microscope equipped with a 60 Plan Apo 1.4 NA oil immersion DIC objective. Images were captured using the Nikon EZ-C1 software (version 2.01). The white bar in all panels represents 20 μm
2
Materials
2.1 Nuclear, Mitochondrial, and Cytosolic Fraction Isolation from Cultured Cells
1. PBS: 137.93 mM NaCl, 2.67 mM KCl, 8.06 mM Na2HPO4, 1.47 mM KH2PO4, pH 7.4. 2. Buffer A: 250 mM sucrose, 20 mM Hepes, 10 mM KCl, 1.5 mM MgCl2, 0.1 mM EDTA, 1 mM EGTA, pH 7.5 (adjusted with KOH). 3. Buffer B: 250 mM sucrose, 10 mM MgCl2. 4. Buffer C: 350 mM sucrose, 0.5 mM MgCl2. 5. Nuclear buffer: 5 mM HEPES, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 26% glycerol (v/v), pH 7.9.
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Western Blotting
1. Protein quantification: Bradford (B6916; Sigma, St. Quentin Fallavier, France) or bicinchoninic acid (BCA) (23227; Thermo Scientific, Rockford, IL, USA) assays. 2. Laemmli buffer: 62.5 mM Tris–HCl at pH 6.8, 25% glycerol, 2% SDS, 0.01% bromophenol blue. Add 5% (about 100 mM) β-mercaptoethanol prior to use (see Note 1). 3. Mini-PROTEAN system with Mini Trans-Blot module, gel cassettes and casting stand, short and spacer plates, combs, external power supply (Bio-Rad, Hercules, CA, USA) and rollers or shakers for incubations. 4. Prestained protein standard (161-0374; Bio-Rad), polyvinylidene difluoride (PVDF) membranes (Millipore, Bedford, MA) and Ponceau S staining solution: 0.1% (w/v) in 5% acetic acid. 5. Buffers: Running buffer (25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3); transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol); and TBS-T (10 mM Tris–HCl at pH 8.0, 150 mM NaCl, 0.1% Tween20). 6. ECF substrate (RPN5785; GE Healthcare, Munich, Germany) and a detection system like VersaDoc (Bio-Rad).
2.3 Immunoprecipitation
1. Protein G PLUS-Agarose (sc-2002, Santa Cruz Biotechnology, Santa Cruz, CA, USA). 2. IP buffer I (50 mM Tris–HCl, pH 7.4, 150 mM NaCl, 1% Na-deoxycholate, 1% NP-40, 1 mM Na3VO4). 3. IP buffer II (50 mM Tris–HCl, pH 7.4, 75 mM NaCl, 0.1% Na-deoxycholate, 0.1% NP-40, 1 mM Na3VO4). 4. IP buffer III (50 mM Tris–HCl, pH 7.4, Na-deoxycholate, 0.05% NP-40, 1 mM Na3VO4).
0.05%
5. Laemmli buffer: 62.5 mM Tris–HCl at pH 6.8, 25% glycerol, 2% SDS, 0.01% bromophenol blue. Add 5% β-mercaptoethanol prior to use. 2.4 Immunocytochemistry
1. Sterile glass coverslips, 6-well plates, lancets, forceps, microscope slides, and a dark humidified chamber. 2. Culture medium and mitochondrial fluorescent probes: MitoTracker Red CMXRos (M7512; Invitrogen, Paisley, UK), MitoTracker Green FM (M7514; Invitrogen), CellLight Mitochondria-GFP, BacMam 2.0 (C10600; Invitrogen), or CellLight Mitochondria-RFP, BacMam 2.0 (C10601; Invitrogen). 3. PBS: 137 mM NaCl in 10 mM phosphate buffer, pH 7.4. 4. 4% formaldehyde in PBS, 0.2% Triton X-100 in PBS, and 1% BSA in PBS.
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5. ProLong Gold Antifade medium with DAPI (P36935; Invitrogen) or without DAPI (P36934; Invitrogen), and Vectashield (preferable) or nail polish (less expensive).
3
Methods
3.1 Nuclear, Mitochondrial, and Cytosolic Fraction Isolation from Cells in Culture
1. Supplement buffer A with 1 μg/ml of leupeptin, antipain, chymostatin, and pepstatin A, 1 mM of DTT, and 100 μM of PMSF. Supplement buffer B and C with 1 mM of DTT and 100 μM of PMSF. Supplement the nuclear buffer with 300 mM of NaCl (high salt helps lyse nuclear membranes and forces DNA into solution). 2. Grow cells on cell culture dishes in an appropriate cell culture medium and perform the desired treatment (see Note 2). 3. After treatment, aspirate or collect (if you are interested in the floating cells) the incubation media. Rinse the adherent cells with 5 ml of PBS and waste it. 4. Harvest adherent cells with 3–5 ml of trypsin. Inhibit the trypsin with 3–5 ml of growth medium containing FBS (see Note 3). 5. Centrifuge between 300 and 400 g for 5 min at 4 C in order to collect all cells without damaging membrane integrity. 6. Aspirate the supernatant and rinse the cell pellet with 2 ml of PBS. Perform again the centrifugation step, discard the supernatant, and resuspend the pellet in 1 ml of complete buffer A. Incubate for 15 min on ice. 7. Homogenize cells in a Potter-Elvehjem homogenizer with a Teflon pestle (30–40 strokes) or, alternatively, pass cell suspension through a 25 G needle 10 times using a 1 ml syringe. This procedure should also be performed on ice. 8. Centrifuge the cellular suspension at 720 g for 5 min at 4 C. Remove the supernatant (containing mitochondrial and cytosolic fractions) and keep it on ice. 9. Resuspend the nuclear pellet again in 1 ml of complete buffer A. Homogenize the pellet again in a Potter-Elvehjem homogenizer with a Teflon pestle or, alternatively, pass through a 25 G needle. Centrifuge again at 720 g for 10 min at 4 C. Discard the supernatant and resuspend the pellet in 0.5 ml of complete buffer B and pour the nuclear suspension on the complete buffer C. Centrifuge at 1430 g for 5 min at 4 C. Remove the supernatant and discard it. Resuspend the nuclear pellet in 50 μl of nuclear buffer supplemented with 300 mM of NaCl. Homogenize the nuclear pellet passing the nuclear suspension through a 27 G needle 10 times using a 0.5 ml syringe. Store at 80 C until further analysis.
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Fig. 3 Flowchart outlining the isolation of nuclear, mitochondrial, and cytosolic fractions from cells in culture. All the material was kept on ice during all the procedure and all the centrifugations were performed at 4 C
10. Centrifuge the supernatant collected in step 8 at 14,000 g during 10 min at 4 C. Resuspend the pellet (mitochondrial fraction) in 50 μl of buffer 1 and store at 80 C until further analysis. 11. Collect the supernatant (containing the cytosolic fraction) and centrifuge in an ultracentrifuge at 100,000 g for 30 min at 4 C. Discard the pellet (containing the membrane fractions) and concentrate the supernatant (containing the cytosolic fraction) by lyophilization or by tangential flow filtration. Store at 80 C until further analysis be performed (see Note 4). In Fig. 3, a flowchart outlining the isolation procedure is presented. 3.2
Western Blotting
3.2.1 SDS-Page
1. Determine the protein content of all fractions by standard procedures such as the Bradford or the BCA method by using 5 μl of aliquot. 2. Mix the samples with the appropriated volume of Laemmli buffer (1:1). 3. Prepare the sodium dodecyl sulfate (SDS) 10% polyacrylamide resolving gel (see Note 5) by mixing 2.5 ml Tris–HCl 1.5 M pH 8.8, 0.1 ml SDS 10%, 50 μl ammonium persulfate 10%, 2.5 ml acrylamide/bisacrylamide (29:1) 40%, 4.9 ml of distilled water (for a 0.75–1.0 mm thick gel). Add 5 μl of TEMED to initiate polymerization. Cast gel within an assembled gel cassette allowing space for stacking gel, gently overlay with water and wait until polymerization.
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4. Prepare the stacking gel (4% polyacrylamide) by mixing 1.25 ml Tris–HCl 0.5 M, pH 6.8, 50 μl SDS 10%, 25 μl ammonium persulfate 10%, 0.5 ml acrylamide/bisacrylamide (29:1) 40%, 3.2 ml of distilled water, and 5 μl of TEMED. Insert a gel comb immediately and wait until polymerization. 5. Denature the samples by boiling at 95–100 C for 5 min. 6. For polyacrylamide gel electrophoresis, we normally use MiniPROTEAN systems from Bio-Rad, but other alternative systems can be used. After complete polymerization, place the gel into the electrophoretic chamber with running buffer, load the volume of sample corresponding to 5–25 μg of protein in each individual well (see Note 6), as well as 6 μl of pre-stained protein standard (e.g., Precision Plus Protein Dual Color Standards from Bio-Rad) into one of the other lanes. 7. Run electrophoresis at constant voltage (100–120 V). Running can be monitored by observing the migration of pre-stained protein standards and bromophenol blue front. Stop running when the bromophenol blue band leaves the lower end of the gel. Keep gels in running buffer until ready to transfer. 3.2.2 Wet/Tank Electrophoretic Transfer
1. Activate the polyvinylidene difluoride (PVDF) membrane in methanol for 15 s. Transfer the membrane from methanol to transfer buffer and incubate on a shaker for at least 5 min. Soak also pads, filter papers, and the gel in transfer buffer before use. 2. Assemble the transfer sandwich in a shallow tray filled with transfer buffer as follows: Black side of the sandwich (cathode), soaked pads, filter paper, gel, PVDF membrane, filter paper, soaked pad, and white side of the sandwich (anode). Avoid bubbles formation between the gel and PVDF membrane. Fill tank apparatus with transfer buffer and run at 350 mA for 90 min with an ice pack or another cooling system (see Note 7). 3. Stain the membrane with Ponceau S for 5 min to check protein transfer. Finally, transfer the membrane to TBS-T to rinse Ponceau staining.
3.2.3 Enzymatic Immunodetection
1. Block the membrane to reduce nonspecific binding with 5% skim milk in TBS-T overnight at 4 C or, alternatively, for 2 h at room temperature (see Notes 8 and 9). 2. Incubate the membrane with the primary antibody at the recommended dilution (see Table 1) in 1% skim milk in TBT-T for 4 h at 4 C or overnight. 3. Wash three times with TBS-T for 5 min at room temperature. 4. Incubate the membrane with the corresponding alkaline phosphatase-conjugated secondary antibody (1:5000) in 1% skim milk in TBT-T for 2 h at 4 C.
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5. Wash three times with TBS-T for 20 min at room temperature. 6. Detect the immunoconjugates using the western blotting ECF substrate (or similar) according to the manufacturer’s protocol. Use fluorescence scanning equipment such as VersaDoc (BioRad) to develop the image and analyze it using the software QuantityOne (Bio-Rad) or Image J (or similar). Please notice that local vs. global background subtraction must be chosen since this will affect the final results. 3.3 Immunoprecipitation
P53 ubiquitination and the physical interaction of p53 with some related proteins such as Bcl-2, Bcl-xL, Bax, and SOD-2 can be evaluated by p53 immunoprecipitation from homogenates and subsequent immunoblot analysis with antibodies against these related proteins. 1. Incubate a volume of extracts corresponding to an equal quantity of protein (e.g., 300 μg of protein) with 5 μl mouse antip53 (2524; Cell Signaling Technology) for 1 h at 4 C under orbital shaking conditions. 2. Add 20 μl of protein G PLUS-Agarose and incubate overnight at 4 C under orbital shaking conditions. 3. Centrifuge 1 min at 16,000 g and discard the supernatant fraction. 4. Add 1 ml of IP buffer I to the pellet and mix the sample on an oscillatory shaker for 20 min at 4 C. 5. Centrifuge for 1 min at 16,000 g and discard the supernatant fraction. 6. Repeat with IP buffer II. 7. Repeat with IP buffer III. 8. Centrifuge 1 min at 16,000 g and discard the supernatant fraction. 9. Add 20 μl of Laemmli buffer and boil for 5 min. 10. Centrifuge again for 1 min at 16,000 g to pellet the agarose beads. 11. Transfer the supernatant fraction to a new tube, subject it to SDS-PAGE and immunoblot analysis as previously described (in Subheading 3.2) using primary antibodies against p53 (sc-6243; Santa Cruz Biotechnology), BCL-2, BCL-xL, BAX, SOD-2, and/or ubiquitin (check Table 1 for references and recommended dilutions).
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1. Seed cells on glass coverslips in 6-well plates and wait 24 h for cell adhesion at 37 C in a 5% CO2 atmosphere.
3.4.1 Cell Seeding for Morphological Studies 3.4.2 Staining with a Mitochondrial-Specific Fluorescent Dye (Option 1)
1. Remove culture medium and incubate cells with 125 nM MitoTracker Red CMXRos (M7512; Invitrogen) in culture medium for 20 min at 37 C. Alternatively, 0.5 μM green-fluorescing MitoTracker Green FM (M7514; Invitrogen) for 30 min at 37 C can be used to label mitochondria (see Note 10). 2. Replace the staining solution with fresh pre-warmed media and subject cells to subsequent processing steps.
3.4.3 Staining with a Mitochondrial-Specific Fluorescent Protein (Option 2)
1. Remove culture medium and add new with 2 μl of CellLight reagent BacMam 2.0 (either GFP-C10600 or RFP-C10601; Invitrogen) per 10,000 cells.
3.4.4 Fixation and Permeabilization
1. Remove the incubation media and fix cells in 4% formaldehyde in PBS for 15 min at 37 C.
2. Mix gently and incubate overnight for at least 16 h.
2. Rinse three times with PBS for 5 min each. 3. Permeabilize cells with 0.2% Triton X-100 in PBS for 10 min. 4. Rinse three times with PBS for 5 min each. 3.4.5 Immunolabeling
1. Block to reduce nonspecific binding with 1% BSA in PBS during 1 h at 4 C (see Note 11). 2. Distribute 100–150 μl of primary antibody (or a mixture of antibodies) at 1:250 in PBS, 1% BSA on each coverslip and incubate 90 min in a humidified chamber at room temperature. 3. Remove the primary antibody (it can be stored in 0.05% sodium azide) and rinse three times with PBS for 5 min. 4. Incubate with the corresponding fluorescence-conjugated secondary antibody (or a mixture of antibodies) at 1:400 in PBS, 1% BSA for 1 h in a dark humidified chamber at 37 C. 5. Remove the secondary antibody and rinse with PBS three times for 5 min each.
3.4.6 Mounting Coverslips
1. Mount coverslips on glass slides using ProLong Gold Antifade medium (P36934; Invitrogen). If nuclear labeling is desired, use ProLong Gold Antifade medium with DAPI (P36935; Invitrogen). 2. Allow to dry overnight, seal with nail polish, and store at 80 C until analysis under a confocal microscope (LSM 510Meta; Zeiss).
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Notes 1. Alternatively to the use of β-mercaptoethanol in western blot sample preparation, 5–10 mM of dithiothreitol (DTT) can be used. Higher concentrations of DTT (65–70 mM) are recommended for proteins with higher amount of disulfide bonds. 2. In order to have enough protein for the different cellular fractions, it is often necessary to start with a large number of cells, especially for large volume cells. An optimization process for cell density is required. 3. If the incubation medium is saved in order to collect floating cells, it can be used to inhibit the trypsin. 4. It is advisable to test the purity of each sample using specific antibodies for each fraction. For example, one strategy involves cross-labeling samples from different fractions with antibodies against a particular histone (nuclear marker), the voltagedependent anion channel (VDAC) or TOM20 (mitochondrial markers) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH, a cytosolic marker). 5. Choose the percentage of the gel to be used according to the molecular weight of proteins of interest. In general: 4–5% gels: >250 kDa; 7.5% gels: 250–120 kDa; 10% gels: 120–40 kDa; 13% gels: 40–15 kDa; 15% gels: