Methods and Protocols

Methods in Molecular Biology 1295

Anton Posch Editor

Proteomic Profiling Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

Proteomic Profiling Methods and Protocols

Edited by

Anton Posch Bio-Rad Laboratories GmbH, Munich, Germany

Editor Anton Posch Bio-Rad Laboratories GmbH Munich, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-2549-0 ISBN 978-1-4939-2550-6 (eBook) DOI 10.1007/978-1-4939-2550-6 Library of Congress Control Number: 2015933382 Springer New York Heidelberg Dordrecht London © Springer Science+Business Media New York 2015 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, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. Printed on acid-free paper Humana Press is a brand of Springer Springer Science+Business Media LLC New York is part of Springer Science+Business Media (www.springer.com)

Preface Sample Preparation Methods and Protein Techniques for Proteomic Profiling This volume is a comprehensive continuation and extension of a book called “2D PAGE: Sample Preparation and Fractionation” which was published in 2008. This book presents the latest developments of the main pillars of protein analysis, namely sample preparation, separation, and characterization. Individual technologies of each pillar combined into complementary and robust workflows render proteomic analysis of complex biological samples even more powerful and are the prerequisite to gain maximum value from biological samples in a single experiment. In this volume, basic but important sample preparation protocols are described again, followed by sophisticated procedures to enrich for specific protein classes and completed by the detailed description of integrated workflows for comprehensive protein analysis and characterization. The authors of the individual chapters are well-known protein biochemists, and all of them have set value to provide a detailed representation of their lab work and to share important tips and tricks for a successful and reproducible employment of their precious protocols in other laboratories. This book is for students of Biochemistry, Biomedicine, Biology, and Genomics and will be an invaluable source for the experienced, practicing scientist, too. Munich, Germany

Anton Posch

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Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Mechanical/Physical Methods of Cell Distribution and Tissue Homogenization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stanley Goldberg 2 Sample Preservation Through Heat Stabilization of Proteins: Principles and Examples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mats Borén 3 Isolating Peripheral Lymphocytes by Density Gradient Centrifugation and Magnetic Cell Sorting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Frederic Brosseron, Katrin Marcus, and Caroline May 4 Investigating the Adipose Tissue Secretome: A Protocol to Generate High-Quality Samples Appropriate for Comprehensive Proteomic Profiling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Simon Göddeke, Jorg Kotzka, and Stefan Lehr 5 Methods for Proteomics-Based Analysis of the Human Muscle Secretome Using an In Vitro Exercise Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mika Scheler, Martin Hrabě de Angelis, Hadi Al-Hasani, Hans-Ulrich Häring, Cora Weigert, and Stefan Lehr 6 Urinary Pellet Sample Preparation for Shotgun Proteomic Analysis of Microbial Infection and Host–Pathogen Interactions . . . . . . . . . . . . . . . . . Yanbao Yu and Rembert Pieper 7 A Protocol for the Parallel Isolation of Intact Mitochondria from Rat Liver, Kidney, Heart, and Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sabine Schulz, Josef Lichtmannegger, Sabine Schmitt, Christin Leitzinger, Carola Eberhagen, Claudia Einer, Julian Kerth, Michaela Aichler, and Hans Zischka 8 Isolation of Mitochondria from Cultured Cells and Liver Tissue Biopsies for Molecular and Biochemical Analyses. . . . . . . . . . . . . . . . . . . . . . . Sabine Schmitt, Carola Eberhagen, Susanne Weber, Michaela Aichler, and Hans Zischka 9 Dynamic Range Compression with ProteoMiner™: Principles and Examples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lei Li 10 Qualitative and Quantitative Proteomic Analysis of Formalin-Fixed Paraffin-Embedded (FFPE) Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Omid Azimzadeh, Michael J. Atkinson, and Soile Tapio

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11 Full-Length Protein Extraction Protocols and Gel-Based Downstream Applications in Formalin-Fixed Tissue Proteomics . . . . . . . . . . . . . . . . . . . . . . Alessandro Tanca, Sergio Uzzau, and Maria Filippa Addis 12 Enrichment of Low-Abundant Protein Targets by Immunoprecipitation Upstream of Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Barbara Kaboord, Suzanne Smith, Bhavin Patel, and Scott Meier 13 Principles of Protein Labeling Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christian Obermaier, Anja Griebel, and Reiner Westermeier 14 Isolation of Extracellular Vesicles for Proteomic Profiling . . . . . . . . . . . . . . . . Dong-Sic Choi and Yong Song Gho 15 A Protocol for Exosome Isolation and Characterization: Evaluation of Ultracentrifugation, Density-Gradient Separation, and Immunoaffinity Capture Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David W. Greening, Rong Xu, Hong Ji, Bow J. Tauro, and Richard J. Simpson 16 Chloroplast Isolation and Affinity Chromatography for Enrichment of Low-Abundant Proteins in Complex Proteomes . . . . . . . . . . . . . . . . . . . . . Roman G. Bayer, Simon Stael, and Markus Teige 17 Depletion of RuBisCO Protein Using the Protamine Sulfate Precipitation Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ravi Gupta and Sun Tae Kim 18 Step-by-Step Preparation of Proteins for Mass Spectrometric Analysis . . . . . . . Thomas Franz and Xinping Li 19 Identification of Protein N-Termini Using TMPP or Dimethyl Labeling and Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jingjing Deng, Guoan Zhang, Fang-Ke Huang, and Thomas A. Neubert 20 Optimization of Cell Lysis and Protein Digestion Protocols for Protein Analysis by LC-MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dominic Winter, Alireza Dehghani, and Hanno Steen 21 Comprehensive Protocol to Simultaneously Study Protein Phosphorylation, Acetylation, and N-Linked Sialylated Glycosylation . . . . . . . Marcella Nunes Melo-Braga, María Ibáñez-Vea, Martin Røssel Larsen, and Katarzyna Kulej 22 Protein Profiling and Phosphoprotein Analysis by Isoelectric Focusing . . . . . . Giuseppina Maccarrone and Michaela D. Filiou 23 Principles and Examples of Gel-Based Approaches for Phosphoprotein Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Birgit Steinberger and Corina Mayrhofer 24 Neutral Phosphate-Affinity SDS-PAGE System for Profiling of Protein Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emiko Kinoshita-Kikuta, Eiji Kinoshita, and Tohru Koike 25 Enrichment and Identification of Bacterial Glycopeptides by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nichollas E. Scott and Stuart J. Cordwell

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26 In-Gel Peptide IEF Sample Preparation for LC/MS Analysis. . . . . . . . . . . . . . Tom Berkelman, Sricharan Bandhakavi, and Aran Paulus 27 Western Blotting Using In-Gel Protein Labeling as a Normalization Control: Stain-Free Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jennifer E. Gilda and Aldrin V. Gomes 28 2-D Western Blotting for Evaluation of Antibodies Developed for Detection of Host Cell Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tom Berkelman, Adriana Harbers, and Sricharan Bandhakavi 29 Free Flow Electrophoresis for Separation of Native Membrane Protein Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lutz Andreas Eichacker, Gerhard Weber, Ute Sukop-Köppel, and Robert Wildgruber 30 Three-Dimensional Electrophoresis for Quantitative Profiling of Complex Proteomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sergio Mauro, Bertrand Colignon, Marc Dieu, Edouard Delaive, and Martine Raes 31 A Bead-Based Multiplex Sandwich Immunoassay to Assess the Abundance and Posttranslational Modification State of β-Catenin . . . . . . . Nicola Groll, Cornelia Sommersdorf, Thomas O. Joos, and Oliver Poetz 32 Identification of SUMO E3 Ligase-Specific Substrates Using the HuProt Human Proteome Microarray . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eric Cox, Ijeoma Uzoma, Catherine Guzzo, Jun Seop Jeong, Michael Matunis, Seth Blackshaw, and Heng Zhu 33 Amyloid-Binding Proteins: Affinity-Based Separation, Proteomic Identification, and Optical Biosensor Validation . . . . . . . . . . . . . . . Alexei Medvedev, Olga Buneeva, Arthur Kopylov, Oksana Gnedenko, Alexis Ivanov, Victor Zgoda, and Alexander A. Makarov 34 Proteomic Profiling by Nanomaterials-Based Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry for High-Resolution Data and Novel Protein Information Directly from Biological Samples . . . . . . Suresh Kumar Kailasa and Hui-Fen Wu Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors MARIA FILIPPA ADDIS • Porto Conte Ricerche, Tramariglio, Alghero(SS), Italy MICHAELA AICHLER • Research Unit Analytical Pathology–Institute of Pathology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany HADI AL-HASANI • Institute of Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Duesseldorf, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany MARTIN HRABĚ DE ANGELIS • Institute of Experimental Genetics, Helmholtz Zentrum München, German Research Center for Environmental Health, Neuherberg, Germany; Center of Life and Food Sciences Weihenstephan, Technische Universität München, Freising-Weihenstephan, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany MICHAEL J. ATKINSON • Institute of Radiation Biology, Helmholtz Zentrum München, German Research Center for Environmental Health, Neuherberg, Germany; Technical University of Munich, Munich, Germany OMID AZIMZADEH • Institute of Radiation Biology, Helmholtz Zentrum München, German Research Center for Environmental Health, Neuherberg, Germany SRICHARAN BANDHAKAVI • diaDexus, South San Francisco, CA, USA ROMAN G. BAYER • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria TOM BERKELMAN • Bio-Rad Laboratories, Hercules, CA, USA SETH BLACKSHAW • Solomon H. Snyder Department of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Institute for Cell Engineering, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Department of Ophthalmology, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Center for High-Throughput Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA MATS BORÉN • Denator AB, Gothenburg, Sweden FREDERIC BROSSERON • Deutsches Zentrum für Neurodegenerative Erkrankungen (DZNE) e.V., Bonn, Germany OLGA BUNEEVA • Institute of Biomedical Chemistry, Moscow, Russia DONG-SIC CHOI • Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea BERTRAND COLIGNON • Département Sciences du Vivant, Centre wallon de Recherches agronomiques, Gembloux, Belgium; URBC-NARILIS, Université de Namur, Namur, Belgium STUART J. CORDWELL • School of Molecular Bioscience, The University of Sidney, Sidney, Australia; Discipline of Pathology, School of Medical Sciences, The University of Sidney, Sidney, Australia; Charles Perkins Centre, The University of Sidney, Sidney, Australia

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Contributors

ERIC COX • Biochemistry, Cellular and Molecular Biology Graduate Program, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Solomon H. Snyder Department of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, MD, USA ALIREZA DEHGHANI • Institute for Biochemistry and Molecular Biology, University of Bonn, Bonn, Germany EDOUARD DELAIVE • URBC-NARILIS, Université de Namur, Namur, Belgium JINGJING DENG • Department of Biochemistry and Molecular Pharmacology, Kimmel Center for Biology and Medicine at the Skirball Institute, New York University School of Medicine, New York, NY, USA MARC DIEU • URBC-NARILIS, Université de Namur, Namur, Belgium CAROLA EBERHAGEN • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany LUTZ ANDREAS EICHACKER • Center of Organelle Research, University of Stavanger, Stavanger, Norway CLAUDIA EINER • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany MICHAELA D. FILIOU • Max Planck Institute of Psychiatry, Munich, Germany THOMAS FRANZ • Max Planck Institute for Biology of Ageing, Cologne, Germany YONG SONG GHO • Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea JENNIFER E. GILDA • Department of Neurobiology, Physiology, and Behavior, University of California, Davis, CA, USA OKSANA GNEDENKO • Institute of Biomedical Chemistry, Moscow, Russia SIMON GÖDDEKE • Institute of Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Leibniz Center for Diabetes Research at Heinrich Heine University, Duesseldorf, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany STANLEY GOLDBERG • Glen Mills Inc., Clifton, NJ, USA ALDRIN V. GOMES • Department of Neurobiology, Physiology, and Behavior, University of California, Davis, CA, USA; Department of Physiology and Membrane Biology, University of California, Davis, CA, USA DAVID W. GREENING • Department of Biochemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia ANJA GRIEBEL • SERVA Electrophoresis GmbH, Heidelberg, Germany NICOLA GROLL • Department of Protein Analytics, NMI Natural and Medical Sciences Institute at the University of Tuebingen, Reutlingen, Germany RAVI GUPTA • Department of Plant Bioscience, Pusan National University, Miryang, Republic of Korea CATHERINE GUZZO • Department of Biochemistry and Molecular Biology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, MD, USA ADRIANA HARBERS • Bio-Rad Laboratories, Hercules, CA, USA HANS-ULRICH HÄRING • Division of Endocrinology, Diabetology, Angiology, Nephrology, Pathobiochemistry and Clinical Chemistry, Department of Internal Medicine, University of Tübingen, Tübingen, Germany; Institute for Diabetes Research and Metabolic Diseases of the Helmholtz Zentrum München at the University of Tübingen, Tübingen, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany

Contributors

xiii

FANG-KE HUANG • Department of Biochemistry and Molecular Pharmacology, Kimmel Center for Biology and Medicine at the Skirball Institute, New York University School of Medicine, New York, NY, USA MARÍA IBÁÑEZ-VEA • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark ALEXIS IVANOV • Institute of Biomedical Chemistry, Moscow, Russia JUN SEOP JEONG • Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, MD, USA HONG JI • Department of Biochemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia THOMAS O. JOOS • Department of Protein Analytics, NMI Natural and Medical Sciences Institute at the University of Tuebingen, Reutlingen, Germany BARBARA KABOORD • Thermo Fisher Scientific, Rockford, IL, USA SURESH K. KAILASA • Department of Applied Chemistry, S. V. National Institute of Technology, Surat, India JULIAN KERTH • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany SUN TAE KIM • Department of Plant Bioscience, Pusan National University, Miryang, Republic of Korea EIJI KINOSHITA • Department of Functional Molecular Science, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan EMIKO KINOSHITA-KIKUTA • Department of Functional Molecular Science, Institute of Biomedical and Health Sciences, Hiroshima University, Hiroshima, Japan TOHRU KOIKE • Department of Functional Molecular Science, Institute of Biomedical and Health Sciences, Hiroshima University, Hiroshima, Japan ARTHUR KOPYLOV • Institute of Biomedical Chemistry, Moscow, Russia JOERG KOTZKA • Institute of Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Leibniz Center for Diabetes Research at Heinrich Heine University, Duesseldorf, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany KATARZYNA KULEJ • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark MARTIN RØSSEL LARSEN • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark STEFAN LEHR • German Center for Diabetes Research (DZD), Neuherberg, Germany; Institute of Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Duesseldorf, Germany CHRISTIN LEITZINGER • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany XINPING LI • Max Planck Institute for Biology of Ageing, Cologne, Germany LEI LI • Bio-Rad Laboratories, Hercules, CA, USA JOSEF LICHTMANNEGGER • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany GIUSEPPINA MACCARRONE • Max Planck Institute of Psychiatry, Munich, Germany ALEXANDER A. MAKAROV • Engelhardt Institute of Molecular Biology of Russian Academy of Sciences, Moscow, Russia

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Contributors

KATRIN MARCUS • Medizinisches Proteom-Center, Ruhr-Universität Bochum, Bochum, Germany MICHAEL MATUNIS • Department of Biochemistry and Molecular Biology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, MD, USA SERGIO MAURO • Département Sciences du Vivant, Centre wallon de Recherches agronomiques, Gembloux, Belgium CAROLINE MAY • Medizinisches Proteom-Center, Ruhr-Universität Bochum, Bochum, Germany CORINA MAYRHOFER • Institute of Animal Breeding and Genetics, University of Veterinary Medicine, Vienna, Austria; Institute of Biotechnology in Animal Production, Department for Agrobiotechnology, University of Natural Resources and Applied Life Sciences Vienna, Tulln, Vienna, Austria ALEXEI MEDVEDEV • Department of Proteomic Research and Mass Spectrometry, Institute of Biomedical Chemistry, Moscow, Russia SCOTT MEIER • Thermo Fisher Scientific, Rockford, IL, USA MARCELLA N. MELO-BRAGA • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark THOMAS A. NEUBERT • Department of Biochemistry and Molecular Pharmacology, Kimmel Center for Biology and Medicine at the Skirball Institute, New York University School of Medicine, New York, NY, USA CHRISTIAN OBERMAIER • SERVA Electrophoresis GmbH, Heidelberg, Germany BHAVIN PATEL • Thermo Fisher Scientific, Rockford, IL, USA ARAN PAULUS • Thermo Fisher Scientific, San Jose, CA, USA REMBERT PIEPER • The J Craig Venter Institute, Rockville, MD, USA OLIVER POETZ • Department of Protein Analytics, NMI Natural and Medical Sciences Institute at the University of Tuebingen, Reutlingen, Germany MARTINE RAES • URBC-NARILIS, Université de Namur, Namur, Belgium MIKA SCHELER • Institute of Experimental Genetics, Helmholtz Zentrum München, German Research Center for Environmental Health, Neuherberg, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany SABINE SCHMITT • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany SABINE SCHULZ • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany NICHOLLAS E. SCOTT • School of Molecular Bioscience, The University of Sidney, Sidney, Australia; Centre for High-Throughput Biology, Department of Biochemistry and Molecular Biology, The University of British Columbia, Vancouver, Canada RICHARD J. SIMPSON • Department of Biochemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia SUZANNE SMITH • Thermo Fisher Scientific, Rockford, IL, USA CORNELIA SOMMERSDORF • Department of Protein Analytics, NMI Natural and Medical Sciences Institute at the University of Tuebingen, Reutlingen, Germany SIMON STAEL • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria; VIB Department of Plant Systems Biology, Gent University, Gent, Belgium HANNO STEEN • Department of Pathology, Boston Children’s Hospital and Harvard Medical School, Boston, MA, USA BIRGIT STEINBERGER • Institute of Animal Breeding and Genetics, University of Veterinary Medicine, Vienna, Austria; Department for Agrobiotechnology, Institute of Biotechnology

Contributors

in Animal Production, University of Natural Resources and Applied Life Sciences Vienna, Tulln, Vienna, Austria UTE SUKOP-KÖPPEL • FFE Service GmbH, Feldkirchen, Germany ALESSANDRO TANCA • Porto Conte Ricerche, Tramariglio, Alghero (SS), Italy SOILE TAPIO • Institute of Radiation Biology, Helmholtz Zentrum München, German Research Center for Environmental Health, Neuherberg, Germany BOW J. TAURO • Department of Biochemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia MARKUS TEIGE • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria; Department of Applied Genetics and Cell Biology, University of Natural Resources and Life Sciences, Vienna, Austria IJEOMA UZOMA • Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, MD, USA SERGIO UZZAU • Porto Conte Ricerche, Tramariglio, Alghero (SS), Italy GERHARD WEBER • FFE Service GmbH, Feldkirchen, Germany SUSANNE WEBER • Institute of Experimental Genetics, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany CORA WEIGERT • Division of Endocrinology, Diabetology, Angiology, Nephrology, Pathobiochemistry and Clinical Chemistry, Department of Internal Medicine, University of Tübingen, Tübingen, Germany; Institute for Diabetes Research and Metabolic Diseases of the Helmholtz Zentrum München at the University of Tübingen, Tübingen, Germany; German Center for Diabetes Research (DZD), Duesseldorf, Germany REINER WESTERMEIER • SERVA Electrophoresis GmbH, Heidelberg, Germany ROBERT WILDGRUBER • FFE Service GmbH, Feldkirchen, Germany DOMINIC WINTER • Institute for Biochemistry and Molecular Biology, University of Bonn, Bonn, Germany; Department of Pathology, Boston Children’s Hospital and Harvard Medical School, Boston, MA, USA HUI-FEN WU • Department of Chemistry, Medical Sciences and Nanotechnology, National Sun Yat-Sen University, Kaohsiung, Taiwan RONG XU • Department of Biochemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia YANBAO YU • The Craig Venter Institute, Rockville, MD, USA VICTOR ZGODA • Institute of Biomedical Chemistry, Moscow, Russia GUOAN ZHANG • Department of Biochemistry and Molecular Pharmacology, Kimmel Center for Biology and Medicine at the Skirball Institute, New York University School of Medicine, New York, NY, USA HENG ZHU • Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, MD, USA; Center for High-Throughput Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA HANS ZISCHKA • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany

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Chapter 1 Mechanical/Physical Methods of Cell Distribution and Tissue Homogenization Stanley Goldberg Abstract This chapter covers the various methods of Mechanical Cell Disruption and Tissue Homogenization that are currently commercially available for processing minute samples (8 M urea, are ideal for extracting samples for 2D PAGE and chromatographic workflows such as LC-MS. On the other hand, SDS, >1 % SDS, based buffers are suitable for SDS-PAGE and western blot workflows. When using SDS buffers, it is recommended to maximize their effectiveness by heating the buffer (>90 °C) before adding it to the sample and after homogenization as instructed below. Other components such as detergents and buffering agents can be added as long as the concentrations of the denaturing agents are not affected.

3.3.2 Buffer to Sample Ratio

To ensure full dispersion of protein–protein complexes formed during cooling after heat stabilization, it is important that sufficient buffer is added to the sample. 1. Weigh samples to be extracted (see Note 12). 2. Calculate the amount of buffer needed to reach a buffer-tosample ratio greater than 10 (>10 μl buffer/mg sample). 3. Add buffer to sample and proceed directly with homogenization.

Heat Stabilization of Proteins 3.3.3 Homogenization of Tissue in 8 M Urea Buffer Using Micro pestle Grinding Followed by Microtip Rod Sonication

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Although heat-stabilized samples will usually homogenize easily, it is very important to ensure that the initial homogenization step is thorough to facilitate the resolubilization of proteins. Soft tissue such as brain can be homogenized just using a microtip sonication rod, firmer tissues, e.g. liver or heart, require physical homogenization, e.g. micro pestle grinding. A combination of physical homogenization followed by microtip sonication rod has been found to be efficient. 1. Place the sample in a Low Bind Eppendorf tube. 2. Crush the sample using a micro pestle until no structure remains (see Note 13). 3. Add sufficient buffer to the sample so the buffer-to-sample ratio is above 10, e.g. >10 μl buffer/mg sample. 4. Continue homogenizing the sample–buffer mixture for at least 2 min until no pieces remain and the mix appears homogeneous. 5. Use a microtip sonication rod (Vibra-Cell, Sonics and Materials) to give the mixture ten bursts of 2 s duration at 40 W setting (see Note 14). 6. Centrifuge homogenate for 10 min at 13,000 × g to pellet insoluble material. 7. Collect supernatant and continue with analysis or store homogenate frozen for later analysis.

3.3.4 Homogenization of Tissue in Hot 1 % SDS Buffer Using Micro pestle Grinding Followed by Heating and Microtip Rod Sonication

1. Heat buffer solution to near boiling, >90 °C, in a water bath. 2. Place the sample in a Low Bind Eppendorf tube. 3. Crush the sample using a micro pestle until no structure remains (see Note 13). 4. Add a sufficient volume of the hot buffer to the sample so the buffer-to-sample ratio is above 10, e.g. >10 μl buffer/mg sample. 5. Continue homogenizing the sample–buffer mixture for at least 2 min until no pieces remain and the mix appears homogeneous. 6. Heat tube containing sample–buffer homogenate in a heating block set to 95 °C for 5 min. 7. Use a microtip sonication rod (Vibra-Cell, Sonics and Materials) give the mixture ten (10) bursts of 2 s duration at 40 W setting (see Note 14). 8. Once repeat steps 6 and 7. 9. Centrifuge homogenate for 10 min at 13,000 × g to pellet insoluble material. 10. Collect supernatant and continue with analysis or store homogenate frozen for later analysis.

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Mats Borén

3.4 Analysis of Extracted HeatStabilized Samples

4

Heat-stabilized samples can be analyzed just as other samples once homogenized and extracted as described above. As long as the buffer is compatible with the intended downstream technique no extra steps are needed. In cases where levels of denaturant interfere with analysis, e.g. in solution antibody-based techniques like ELISA, dilution can be useful. Removal of denaturants by buffer exchange, e.g. size exclusion chromatography, can be problematic as denatured proteins have lower solubility in non-denaturing buffers. This can cause precipitation of proteins as the denaturant is removed and should be done with caution.

Notes 1. Denaturing agents of buffers should be >1 % SDS or >8 M urea. In addition to denaturing agents other components can be added, e.g. detergents and salts, depending on the needs of the specific work flow, the SDS and/or urea contents may however not be lowered. 2. Use dust mask to avoid getting powdered SDS into the lungs during weighing. It is also a good idea to make a 10 % SDS stock solution to use for further dilutions to minimize the number of times powdered SDS have to be handled. 3. As urea dissolves the solution becomes cold. Do not heat to speed up solubilization as this may create isocyanic acid which will cause protein carbamylation in later steps. Isocyanic acid can also form naturally as the urea buffer ages. To avoid this, always prepare urea containing extraction buffers fresh prior to use. 4. Both Fresh methods, Quick and Structural preserve, will inactivate enzymes in the sample. The Quick method will compress the sample to make it thinner, giving a faster heating throughout the sample. The Structure preserve on the other hand use minimal compression to preserve sample structure. The sample is thus thicker and stabilization takes longer time. 5. The Post mortem clock is basically a timer keeping track of the time between sacrifice and stabilization. It is important to try to keep this time as short and comparative between samples as possible. The use of the timer helps to focus attention on the dissection and avoids unnecessary delays. As dissection is an art which needs practice it is quite common for the dissection time to decrease by ~50 % between the first and the fifth animal before leveling out. It is thus good practice to either start with control animals not part of the study or a group where time is not as important.

Heat Stabilization of Proteins

31

6. Cervical dislocation or a scientific Guillotine are examples of preferred sacrificial techniques. 7. Card can remain open without support if first folded 180°. To minimize air in cavity after closing it, push the upper foil inwards toward the sample prior to placing the sample in the cavity. 8. Although the heat stabilization will inactivate enzymes, other protein-modifying processes, e.g. oxidation, are still active and could induce protein changes in the sample. Stable levels of phosphorylations have been shown for up to 24 h in room temperature after heat stabilization but it is not recommended to keep stabilized samples at ambient temperatures for more than a few hours at most [9]. 9. Both Frozen methods, Quick and Structural preserve, will inactivate enzymes in the sample. The Quick method will compress the sample to make it thinner, giving a faster heating throughout the sample. The Structure preserve on the other hand use minimal compression to preserve sample structure. The sample is thus thicker and stabilization takes longer time. 10. The algorithms controlling treatment time for the Frozen methods are for treating samples between −78 °C and +20 °C. If samples are transported in liquid nitrogen, −196 °C, they need to be equilibrated either at −20 °C or on dry ice so they are not too cold when treated as this will result in under treatment and residual enzymatic activity. 11. If the Maintainor Tissue cards have been cooled on dry ice longer than 1 min, hold the card between thumb and index finger just over the orange seal during transfer to thaw it slightly. If the orange seal has frozen solid the needle can have difficulty penetrating the septum. 12. This is greatly facilitated if samples are collected in pre-weighted containers. Weigh samples one by one transferring them quickly from dry ice to the scale and read the weight as soon as reasonable stable. Moisture in the air will start to deposit on the sample container during weighing so the scale will not fix on a value. 13. Pre-crushing the sample prior to adding buffer makes micropestle homogenization much easier. If the buffer is added first it can be quite difficult to hunt the small slippery pieces swirling around in the buffer. 14. Have sonicator tip in buffer while active and do not dip it up and down to avoid foaming. This is especially relevant with SDS-based buffers.

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References 1. Sköld K, Alm H, Scholz B (2013) The impact of biosampling procedures on molecular data interpretation. Mol Cell Proteomics 12(6): 1489–1501 2. Stingl C, Söderquist M, Karlsson O et al (2014) Uncovering effects of ex vivo protease activity during proteomics and peptidomics sample extraction in rat brain tissue by oxygen-18 labeling. J Proteome Res 13(6):2807–2817 3. Svensson M, Borén M, Sköld K et al (2009) Heat stabilization of the tissue proteome: a new technology for improved proteomics. J Proteome Res 8(2):974–981 4. Kultima K, Sköld K, Borén M (2011) Biomarkers of disease and post-mortem changes—heat stabilization, a necessary tool for measurement of protein regulation. J Proteomics 75(1): 145–159 5. Ahmed MM, Gardiner KJ (2011) Preserving protein profiles in tissue samples: differing out-

6.

7.

8.

9.

comes with and without heat stabilization. J Neurosci Methods 196(1):99–106 Robinson AA, Westbrook JA, English JA et al (2009) Assessing the use of thermal treatment to preserve the intact proteomes of postmortem heart and brain tissue. Proteomics 9(19):4433–4444 Smejkal GB, Rivas-Morello C, Chang JH et al (2011) Thermal stabilization of tissues and the preservation of protein phosphorylation states for two-dimensional gel electrophoresis. Electrophoresis 32(16):2206–2215 Li X, Friedman BA, Roh MS, Jope RS (2005) Anesthesia and post-mortem interval profoundly influence the regulatory serine phosphorylation of glycogen synthase kinase-3 in mouse brain. J Neurochem 92:701–704 Borén M (2011) Methodology and technology for stabilization of specific states of signal transduction proteins. Methods Mol Biol 717:91–100

Chapter 3 Isolating Peripheral Lymphocytes by Density Gradient Centrifugation and Magnetic Cell Sorting Frederic Brosseron, Katrin Marcus, and Caroline May Abstract Combining density gradient centrifugation with magnetic cell sorting provides a powerful tool to isolate blood cells with high reproducibility, yield, and purity. It also allows for subsequent separation of multiple cell types, resulting in the possibility to analyze different purified fractions from one donor’s sample. The centrifugation step divides whole blood into peripheral blood mononuclear cells (PBMC), erythrocytes, and platelet-rich plasma. In the following, lymphocyte subtypes can be consecutively isolated from the PBMC fraction. This chapter describes enrichment of erythrocytes, CD14-positive monocytes and CD3-positive T lymphocytes. Alternatively, other cell types can be targeted by using magnetic beads specific for the desired subpopulation. Key words Peripheral blood mononuclear cells (PBMC), Magnetic-activated cell sorting (MACS™), Fluorescence-activated cell sorting (FACS)

1

Introduction Isolation of blood cells for research purposes should ensure optimal experimental conditions by providing maximum specificity, purity, yield, speed, and reproducibility [1, 2]. Today, numerous methods for enrichment of different cell types exist, the most common being density gradient centrifugation and several variants of antibody-based immunoaffinity methods [3, 4]. Density gradients are typically used to fractionate full blood into platelet-rich plasma, erythrocytes, and peripheral blood mononuclear cells (PBMCs) composed of lymphocytes, monocytes, and macrophages [5, 6]. Cell (sub)types can be efficiently obtained by use of immunoaffinity purifications [4]: These methods make use of antibodies targeting characteristic proteins like cell surface receptors, for example to attach labels for instrumental supported isolations like fluorescence-activated cell sorting (FACS) [7]. Likewise, antibodies are immobilized on supporting material with properties suited for enrichment methods (e.g., magnetic cell sorting, MACS™).

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 1295, DOI 10.1007/978-1-4939-2550-6_3, © Springer Science+Business Media New York 2015

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The MACS™ technology has been frequently used and described by us and others in terms of efficiency and yield [8, 9]. In this chapter, we describe the use of density gradient centrifugation combined with MACS™ sorting for subsequent isolation of erythrocytes, monocytes, and T lymphocytes from human full blood (Fig. 1). This strategy offers multiple advantages: It is possible to obtain multiple cell types from one blood donation, enabling researchers to investigate multiple targets and make optimal use of the donor sample. The procedure can be easily adapted to other cell types by choice of the respective MACS™ beads, which are commercially available. Furthermore, magnetic cell sorting is of ease of use and therefore also suited for users inexperienced with blood cell isolation.

Diluted whole blood Pancoll

Density gradient centrifugation

Platelet-rich plasma PBMC’s Pancoll Erythrocytes

PBMC’s

MACS™

CD14+ MACS™

CD14+ Monocytes

Erythrocytes

CD3+ MACS™

CD3+ T-Lymphocytes

FACS

Fig. 1 Overview of workflow. He isolation procedure begins with a density gradient centrifugation which fractionates whole blood into erythrocytes, PBMCs, and platelet-rich plasma. Fractions of interest—e.g., erythrocytes—can be preserved for analysis directly after the gradient. The PBMC fraction can be further subdivided by subsequent cell type-specific magnetic cell sorting, as shown in this protocol for CD14-positive monocytes and CD3-positive T lymphocytes

Lymphocyte Isolation by Magnetic Cell Sorting

2

35

Materials

2.1 Technical Equipment

1. Laboratory area for handling of human biomaterial according to local regulations.

2.1.1 General

2. Laminar flow hood, HEPA-filtered and UV-sterilized. 3. Containment for biofluid waste. 4. Water squirt bottle. 5. Disinfectant. 6. Ice box. 7. 4 °C Freezer. 8. −80 °C Freezer.

2.1.2 Blood Sampling

1. 20 mL syringes. 2. Butterfly or similar needle. 3. Adapters to connect needle to syringes.

2.1.3 Density Gradient

1. Centrifuge for 50 mL tubes, swing bucket rotor, adjustable acceleration and breaking, cooling option. 2. Sterile serological pipettes. 3. Pipetting device or Peleus ball for serological pipettes. 4. Sterile 15 and 50 mL conical centrifuge tubes. 5. Stand for centrifuge tubes. 6. 5 mL Plastic Pasteur pipette. 7. Automatic cell counter or microscope with Neumann chamber.

2.1.4 MACS Isolation

1. Magnetic stirring unit. 2. 500 mL cell culture bottles. 3. Sterile filtering device. 4. Precision scales. 5. pH-meter. 6. MACS magnet (Mini MACS and Midi MACS or Vario MACS). 7. MACS™ MS-columns. 8. MACS™ LS-Columns. 9. Cell filter fitting to MACS™ columns. 10. Sterile serological pipettes. 11. Pipetting device or Peleus ball for serological pipettes. 12. Centrifuge for 15 and 50 mL tubes, swing bucket rotor, adjustable acceleration and breaking, cooling option. 13. Sterile 15 and 50 mL conical centrifuge tubes. 14. Stand for centrifuge tubes.

36


15. Micropipettes. 16. Sterile micropipette tips. 17. Automatic cell counter or microscope with Neumann chamber. 2.1.5 Washing of Purified Monocytes and T Lymphocytes

1. Centrifuge for 15 and 50 mL tubes, swing bucket rotor, adjustable acceleration and breaking, cooling option. 2. Sterile serological pipettes. 3. Pipetting device or Peleus ball for serological pipettes. 4. Sterile 15 and 50 mL conical centrifuge tubes. 5. Stand for centrifuge tubes. 6. Table centrifuge for 1.5 mL reaction tubes, cooling option. 7. 1.5 mL sterile polypropylene reaction tubes. 8. Stand for reaction tubes. 9. Micropipettes. 10. Sterile micropipette tips.

2.1.6 Quality Control Using FACS

1. Automatic cell counter or microscope with Neumann chamber. 2. Micropipettes. 3. Sterile micropipette tips. 4. Vortex. 5. Flow cytometer. 6. FACS-vials fitting to flow cytometer.

2.2 Buffers and Solutions

1. Phosphate buffered solution (PBS), sterile, calcium-free, magnesium-free (PBS−/−).

2.2.1 General 2.2.2 Blood Sampling

1. Heparin 25,000 solution.

2.2.3 Density Gradient

1. Hanks buffered salt solution (HBSS). 2. Pancoll® human, mentioned in the following as Pancoll®.

2.2.4 MACS Isolation

1. MACS™ buffer: 0.5 % BSA, 0.06 % EDTA in PBS−/−. Adjust to pH 7.2 using NaOH and HCl, sterile filter and store at 4 °C for a maximum of 1 week (see Note 1). 2. MACS™ CD14 microbeads. 3. MACS™ CD3 microbeads.

2.2.5 Quality Control Using FACS

1. Anti-CD14-FITC. 2. Anti-CD3-PE. 3. Anti-CD19 FITC. 4. Anti-CD56 PE.


3 3.1

37

Methods Blood Sampling

1. Blood withdrawal is to be performed by authorized medical personal only according to local legislative regulations. 2. Prepare 4× 20 mL syringes by placing 50 μl Heparin 25,000 as anticoagulant into the inlet of the syringe before blood sampling (see Note 2). 3. Fill each syringe by venipuncture of cubital vein (see Note 3). 4. Slowly invert syringes three times to ensure homogenous mixing of blood and anticoagulant. 5. Proceed to density gradient immediately.

3.2

Density Gradient

1. Let Pancoll® human adjust to room temperature for 1 h (see Note 4). 2. Dilute 20 mL venous blood 1:1 with HBSS in a 50 mL tube. Invert three times to ensure homogenous mixing (see Note 5). 3. Place 10 mL Pancoll® in a 50 mL centrifuge tube. 4. Layer diluted venous blood slowly and carefully on top of the Pancoll®, using an automatic pipetting device on the lowest setting or a Peleus ball (see Fig. 2). 5. Centrifuge for 20 min, RT, 1,250 × g using a swing bucket rotor, with settings for acceleration and deceleration as low as possible (see Note 6).

Diluted blood Pancoll® Fig. 2 Preparation of density gradient. To prepare the density gradient, begin by placing a small volume of diluted blood as slow as possible on top of the Pancoll® at the edge of the centrifuge tube. Stabilize the pipette by pressing it at the edges of the centrifuge tube. Wait until the blood has distributed over the Pancoll®, then slowly move the pipette up while continuously letting the diluted blood flow out. Avoid too fast outlet of the blood as this will cause disturbances in the gradient, resulting in poor performance

38


6. The gradient should result in four clearly separated phases, listed in order of increasing density: 1st supernatant (yellowish, remaining serum constituents and platelets), 2nd PBMCs (grey phase), 3rd Pancoll® (clear), and 4th erythrocytes (dark red) at the tube bottom (see Fig. 1). 7. Perform all following steps consequently on ice or at 4 °C with precooled buffers and reagents (4 °C). 8. Prepare 2× 50 mL centrifuge tubes with 10 mL cold PBS−/− each. 9. Remove supernatant of gradient using a serological pipette to 1 cm above the PBMC phase. 10. Collect the grayish PBMCs by using a plastic Pasteur pipette, starting at the edge of the tube and later moving across the plane of the PBMC layer. Place PBMCs in the prepared 50 mL tubes containing the PBS−/− (see Subheading 3.4 and Note 7). 11. Remove remaining Pancoll® using a serologic pipette. 12. Use remaining erythrocyte suspension for washing (see Subheading 3.3). 3.3 Preparation of Erythrocytes

1. Fill up tube from step 12 in Subheading 3.3 to 50 mL using PBS−/−. 2. Centrifuge the erythrocytes for 8 min, 4 °C, 470 × g with medium acceleration and deceleration to remove remaining contaminants from the erythrocytes (see Note 8). 3. Discard the supernatant and the top 1–2 mL of the erythrocyte fraction to ensure complete removal of contaminants. 4. Repeat steps 2 and 3 with the erythrocyte fraction. 5. Collect purified erythrocytes in one centrifuge tube and store at −80 °C until usage.

3.4 Washing of PBMCs

1. Fill centrifuge tube from step 3.3. 10 to 50 mL with PBS−/− and invert 3 times to wash the PBMC fraction. 2. Centrifuge the PMBCs for 8 min, 4 °C, 470 × g to remove remaining contaminants (see Note 9). 3. Discard supernatant and resuspend the PBMCs in 1 mL PBS−/−. 4. Unite PBMCs from two tubes into one fresh 50 mL centrifugation tube, fill up to 50 mL with PBS−/− and repeat step 2. 5. Discard supernatant and resuspend PBMCs from all remaining tubes in 1 mL PBS. 6. Take up an aliquot for cell counting. Use a light microscope combined with a Neubauer counting chamber or an automated cell counter according to the manufacturer’s instructions. 7. Preserve 3 × 200,000 cells for FACS analysis of the PBMC fraction (see Note 10).


39

8. After cell counting, wash PBMCs a third time in PBS−/− and remove supernatant. 9. Use PBMC pellet for MACS™ cell isolation (see Subheading 3.6). 3.5 Isolation of CD14+ Monocytes and CD3+ T Lymphocytes

1. Resuspend the PBMC pellet from step 9 in Subheading 3.5 in MACS™ buffer in a 15 mL centrifugation tube according to the manufacturer’s instructions. 2. Add CD14+ MACS™ beads according to the manufacturer’s instructions and vortex. 3. Incubate 4 °C for 15 min. 4. Fill tube to 15 mL with MACS™ buffer, invert 3 times and centrifuge for 10 min, 4 °C, 300 × g. 5. Discard supernatant and resuspend the cell pellet in MACS™ buffer according to the manufacturer’s instructions. 6. Purify CD14+ monocytes using a MACS™ MS column in a Mini MACS™ magnet, cell filter attached, according to the manufacturer’s instructions. 7. Collect both flow-through (contains CD14 depleted PBMCs) and elution fraction (contains CD14+ monocytes) in a separate 15 mL centrifugation tube. 8. Wash elution fraction (see Subheading 3.7). 9. Fill flow-through to 15 mL with MACS buffer and wash as in step 13. 10. Resuspend PBMCs from flow-through in MACS buffer for CD3+ sorting according to the manufacturer’s instructions. 11. Add CD3+ MACS™ beads according to the manufacturer’s instructions and vortex. 12. Incubate 4 °C for 15 min. 13. Fill tube to 15 mL with MACS buffer, invert 3 times and centrifuge for 10 min, 4 °C, 300 × g. 14. Discard supernatant and resuspend the cell pellet in MACS buffer according to the manufacturer’s instructions. 15. Purify CD3+ T lymphocytes using a MACS LS column in a Midi MACS magnet, cell filter attached, according to the manufacturer’s instructions. 16. Collect elution fraction (contains CD3+ T lymphocytes) in a 15 mL centrifugation tube. 17. Wash elution fraction (see Subheading 3.7).

3.6 Washing of Purified Monocytes and T Lymphocytes

1. Resuspend cell pellets from step 7 in Subheading 3.6 or step 16 in Subheading 3.6 in 1 mL PBS−/−. 2. Fill to 15 mL with PBS−/−, invert 3 times, and centrifuge for 10 min, 4 °C, 300 × g.

40


3. Discard supernatant, resuspend in 1 mL PBS−/− and count cells. 4. Preserve 3× 200,000 cells for FACS analysis of the purified cells. 5. Transfer the resuspended cells into a 1.5 mL reaction tube and fill with PBS−/−. 6. Centrifuge the CD14+ cells for 5 min, at 4 °C and 1,000 × g. 7. Remove supernatant completely and store the dry cell pellet at −80 °C until usage. 3.7 Quality Control Using FACS

1. For each isolated fraction (PBMCs, Monocytes, T lymphocytes), label 3 FACS vials as follows: (a) Non-labeled control (b) CD14 FITC + CD3 PE (c) CD19 FITC + CD56 PE 2. Pipette 200,000 cells of the respective isolated fraction in each vial. 3. Incubate cells with FACS antibodies according to the vial labels and manufacturer’s instructions. 4. Measure on a flow cytometer according to the manufacturer’s instructions (see Note 11). 5. Determine composition of fractions by comparing ratios of CD14+ (monocytes), CD3+ (T lymphocytes), CD19+ (B lymphocytes), and CD56+ (NK cells). The isolation of monocytes and T lymphocytes by MACS should provide purities > 85 % (Fig. 3).

4

Notes 1. The MACS™ buffer is best prepared on the day before the cell isolation. 2. The described protocol was originally designed for large yields for proteomics experiments without further culture of the cells and is therefore based on a large blood donation of 80 mL [9]. The procedure is adaptable to any other volume by proportionally adjusting volumes of Pancoll® and MACS™ beads. Magnetic cell sorting can also be performed directly from full blood, but the density gradient will help to pre-purify and concentrate the PBMCs in a small volume for MACS™ isolation. 3. The appropriate blood sampling system/adapters should be chosen by authorized and experienced medical personal. 4. Cold Pancoll® has a different density than warm Pancoll® which can cause inefficient separation in density gradient. Remove Pancoll® from freezer before blood sampling is performed to ensure that there is no time delay between sampling and gradient.

41

Lymphocyte Isolation by Magnetic Cell Sorting MACS CD14+ Fraction

PBMC-Fraction CD3+

FL2-Height

FL2-Height

CD3+

102 101

unlabeled cells

0.2%

59.1% 103

28.3%

12.4% CD14+

100 100

101

102

Unlabeled cells

4.0%

0.4%

3.1%

92.5%

103 102 101

CD14+

100 100

103 104

101

102

103

104

FL1-Height

FL1-Height

MACS CD3+ Fraction

FL2-Height

CD3+

94.9%

0.4%

2.4%

2.3%

103 102 101

CD14+ Unlabeled cells

100 100

101

102

103 104

FL1-Height

Fig. 3 Purity control by FACS. Lymphocyte fractions analyzed by FACS. In fractions isolated from PBMCs by MACS, proportions of the selected cells are drastically increased: In this example, the PBMC fractions consist of 46 % of CD3+ cells (T lymphocytes) and 3 % of CD14+ cells (monocytes). By MACS sorting, these proportions are increased to 88 % (CD14+ fraction) and 95 % (CD3+ fraction)

5. This step is necessary to adjust the density of the blood for the gradient. 6. Fixed angle rotors and strong acceleration or deceleration will cause inefficient separation in gradient. 7. When collecting the PBMCs, include parts of the supernatant and Pancoll® to ensure complete harvest of the PBMCs. The following washing and magnetic cell isolation will remove the contaminants. 8. Erythrocytes will not form a pellet, but a concentrated suspension. 9. This step can be performed together with step 2 in Subheading 3.4. 10. The necessary amount of cells for FACS analysis depends on the used instrument and measurement protocols. 11. FACS analysis should be performed in a lab with high experience in analysis of mononuclear cells.

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Acknowledgement This work was supported by P.U.R.E. (Protein Unit for Research in Europe), a project of Nordrhein-Westfalen, a federal state of Germany, the Bundesministerium für Bildung und Forschung (NGFNplus, FZ 01GS08143), Germany, and the German Center for Neurodegenerative Diseases (DZNE e.V.) within the Helmholtz Association. References 1. Blonder J, Issaq HJ, Veenstra TD (2011) Proteomic biomarker discovery: it’s more than just mass spectrometry. Electrophoresis 32: 1541–1548 2. Mallick P, Kuster B (2010) Proteomics: a pragmatic perspective. Nat Biotechnol 28:695–709 3. Bennett S, Breit SN (1994) Variables in the isolation and culture of human monocytes that are of particular relevance to studies of HIV. J Leukoc Biol 56:236–240 4. Dainiak MB, Kumar A, Galaev IY et al (2007) Methods in cell separations. Adv Biochem Eng Biotechnol 106:1–18 5. Syrovets T, Tippler B, Rieks M et al (1997) Plasmin is a potent and specific chemoattractant for human peripheral monocytes acting via a cyclic guanosine monophosphate-dependent pathway. Blood 89:4574–4583

6. de Almeida MC, Silva AC, Barral A et al (2000) A simple method for human peripheral blood monocyte isolation. Mem Inst Oswaldo Cruz 95:221–223 7. Gorelik L, Kauth M, Gehlhar K et al (2008) Modulation of dendritic cell function by cowshed dust extract. Innate Immun 14: 345–355 8. Mayer A, Lee S, Lendlein A et al (2011) Efficacy of CD14(+) blood monocytes/macrophages isolation: positive versus negative MACS protocol. Clin Hemorheol Microcirc 48:57–63 9. Brosseron F, May C, Schoenebeck B et al (2012) Stepwise isolation of human peripheral erythrocytes, T lymphocytes, and monocytes for blood cell proteomics. Proteomics Clin Appl 6:497–501

Chapter 4 Investigating the Adipose Tissue Secretome: A Protocol to Generate High-Quality Samples Appropriate for Comprehensive Proteomic Profiling Simon Göddeke, Jorg Kotzka, and Stefan Lehr Abstract In this chapter, we describe in detail how to prepare a sample containing the complete entity of secretion products from murine primary adipocytes, which are suitable for comprehensive and sensitive secretome analysis. The underlying protocol should be seen as a starting point guiding through critical steps of the complex workflow in order to approximate to the real secretome in the context of different sample types used for the diverse research questions the protocol has to be carefully adjusted. Key words Secretome analysis, Adipokines, Murine adipose tissue, Pre-adipocytes

1

Introduction Obesity, which is favored by imbalanced energy supply and energy consumption coupled with a sedentary lifestyle is considered as an epidemic disease and represents a burden for almost all societies [1]. Therefore, overweight today is the major risk factor for developing various metabolic complications such as insulin resistance, type 2 diabetes, non-alcoholic liver disease, and cardiovascular diseases [2–6]. Over the last decade we have learned that adipose tissue, besides its predominant role within energy homeostasis also represents a major endocrine organ, releasing a wide variety of signaling and mediator proteins. These so-called adipokines seem to be causally involved in the development of a wide variety of metabolic diseases. Over the last years, several attempts have been made to characterize the adipokines by utilizing diverse proteomic profiling approaches. This has led to a catalog of adipokines comprising several hundred potentially secreted peptides and proteins [7]. Nevertheless, the limited overlap of identified adipokines in published studies impressively


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Simon Göddeke et al.

illustrates that the analysis of tissue-specific secretomes is an extreme challenging business. Within the complex workflow including sample preparation, concentration, proteomic profiling and extensive bioinformatics, the most striking point is to discriminate the real secretome from contaminating proteins. Therefore, a meaningful characterization inalienable starts with high-quality samples and requires a reproducible processing as crucial points for successful identification of secreted proteins. Particularly during sample collection and culturing processes considerable contaminations could occur from different sources, e.g. release of cellular proteins due to cell damage or contamination of high-abundant proteins derived from culture, i.e. FCS. Due to the fact that the expected concentrations of adipokines are low (pg to ng/ml), artificial proteins have the potential to shift the dynamic range of the secretome sample dramatically. In the light of these possible pitfalls, a careful evaluation of sample preparation and culture setup to minimize contamination are the most crucial steps in order to obtain relevant secretome data [7, 8].

2

Materials

2.1 General Hardware and Consumables

1. Incubator (37 °C; 5 % CO2; 95 % humidity). 2. Sterile cell culture bench. 3. Centrifuge. 4. Ultracentrifuge. 5. Laboratory scissors (double sharped); 145 mm. 6. Scalpel grip Nr. 4 L. 7. Scalpel blades Nr. 22. 8. Forceps. 9. 6-Well-culture plates. 10. Polypropylene (PP) centrifugation tubes, 15 ml. 11. Polypropylene (PP) centrifugation tubes, 50 ml. 12. Sterile serological pipettes, 10 ml. 13. Glass funnel, pasteur pipettes. 14. Incubation rotator. 15. Steam-sanitizer. 16. Special accuracy scale.

2.2

Tissue Biopsies

2.3 Isolation of Pre-adipocytes

1. Hank’s Balanced Salt Solution (HBSS) w/o Calcium and Magnesium (pH 7.4). 1. Cell Strainer, 70 μm, Nylon (BD Falcon). 2. Mesh, 150 μm.

Secretome Analysis of Adipose Tissue

45

3. Mesh, 75 μm. 4. Syringe filters, 25 μm. 5. Special accuracy scale. 6. Collagenase (NB8, Clostridium Histolyticum). 7. Phosphate buffered saline (PBS) w/o Calcium and Magnesium, sterile. 8. Collagenase solution (750 U collagenase/g adipose tissue): 250 U/ml in HBSS (pH 7.4) (see Note 1) supplemented with 3 mM CaCl2 (see Note 2) and 4 mM glucose. Sterile filtration (10 ml syringe), G26 cannula (1½″), sterile syringe filters (0.2 μm, 33 mm). 9. HBSS to dilute the collagenase: add CaCl2 (3 mM). 10. Erythrocyte lysis buffer: 8.29 g/l NH4Cl, 0.99 g/l K2HPO4, 0.04 g/l EDTA, pH 7.3 sterile filtration. 11. Basal medium: DMEM/F-12 (17.5 mM glucose), 1.25 g/l NaHCO3, 16 mg/l Biotin, 8 mg/l Calcium-D-pantothenate, pH 7.3, sterile filtration. Before use add 5 ml/l Gentamycin. 12. Basal medium w/o phenol-red: DMEM, w/o phenol red, w/o FCS, 10 pmol/ml insulin. Before use add 5 ml/l Gentamycin. 2.4 Differentiation of Pre-adipocytes

1. Vacuum filtration unit, 500 ml (0.22 μm). 2. Ethylendiaminetetraacetic acid (EDTA) solution pH 8.0 (1 mg/ml). 3. Dulbecco’s Modified Eagle’s Medium/F-12 (DMEM/F-12, 17.5 mM glucose). 4. Phosphate buffered saline (PBS) w/o Calcium and Magnesium, sterile. 5. Cultivation medium: Basalmedium (Subheading 2.3, item 12), 10 % FCS, 5 ml/l Gentamycin. 6. Troglitazon (TZO) stock solution: Solve 5 mg in 1 ml Dimethylsulfoxide (DMSO). 7. Differentiation medium #1 (for 50 ml) (see Note 3): 50 ml basal medium (Subheading 2.3, item 12), 5 μl TZO stock solution (see Note 4), 25 μl Apo-Transferrin stock solution, 100 μl Hydrocortisol stock solution, 5 μl T3 stock solution, 150 μl insulin stock solution. 8. Differentiation medium #2: Differentiation medium #1 w/o TZO, supplement with 10 % FCS. 9. Apo-Transferrin stock solution: Solve 50 mg in ml ddH2O. 10. Hydrocortisol stock solution (50 μg/ml): Solve 1 mg in 1 ml EtOHabs and dilute in basal medium w/o phenol-red.

46


11. Trijodthyronin (T3) stock solution (20 μg/ml): Solve 5 mg in 1 ml 1 N NaOH and dilute in ddH2O. 12. Insulin stock solution (=10−4 M): Solve 5 mg in 860 μl 0.01 N HCl and dilute in 10 ml ddH2O. 2.5 Quality Control of Differentiation by Oil Red O Protocol

1. Oil Red O stock solution: Mix stock solution in 15 ml PP centrifugation tubes with ddH2O (6:4, v/v) and incubate for at least 2 h at room temperature in the dark. Filtrate the solution before use and centrifuge for 10 min at 3,200 × g. 2. Hemalaun solution: Dissolve Hematoxylin in ddH2O (1 mg/ml) and add sodium iodate (0.2 mg/ml) and potassium sulfate (50 mg/ml) while agitating (see Note 5). After mixing add chloride-hydrate (50 mg/ml) and crystalline citric acid (1 mg/ml) (see Note 6). 3. Fixation solution: Mix 15 ml Picric acid with 5 ml Formol (37 %) and 1 ml acetic acid (100 %). 4. Oil Red O stock solution: 0.3 g Oil Red O solve in 100 ml isoproanol (99 %) (see Note 7).

2.6 Collecting Secreted Peptides/ Proteins

1. Mesh, 50 μm. 2. Ethylendiaminetetraacetic acid (EDTA) solution pH 8.0 (1 mg/ml). 3. Phosphate buffered saline (PBS) w/o Calcium and Magnesium, sterile.

2.7 Concentration via Centrifugal Filter Concentrator

3

1. Centrifugal filter concentrator (Amicon Ultra 15, 3 kDa).

Methods In this chapter we describe in detail how to prepare a specimen, containing the complete entity of secretion products from murine primary adipocytes, which are suitable for comprehensive and sensitive secretome analysis. The underlying protocol should be seen as a starting point guiding through critical steps of the complex workflow in order to approximate to the real secretome in the context of different sample types used for the diverse research questions the protocol has to be carefully adjusted (Fig. 1).

3.1

Tissue Biopsies

1. On the day of surgery, after a 6 h fast, mice are sacrificed and adipose tissue specimens are obtained from the subcutaneous, visceral or brown depots. 2. Harvest the tissue as a whole part, not in small pieces. 3. Tissue specimens were immediately transferred in HBSS and transported on ice to the laboratory (see Note 8).


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Fig. 1 Workflow scheme to generate secretome sample from differentiated mouse adipocytes

3.2 Isolation of Pre-adipocytes (Figs. 2, 3, and 4)

1. Dissect tissue from fibrous material and visible blood vessels and cut into fragments of ∼5–10 mg (see Note 9). 2. Transfer the mashy tissue in a 50 ml tube and add collagenase mixture (see Note 10). 3. Incubate fat tissue fragments with vigorous (250 cycles/min) for at least 30 min at 37 °C.

shaking

4. Centrifugation: 5 min, 240 × g, RT (see Note 11). 5. Re-suspend the pellet in HBSS (1:10, v/v). 6. The supernatant, more precisely the floating mature adipocytes, is harvested and transferred to another 50 ml tube with a Pasteur pipette (see Note 12). 7. Stop collagenase reaction by adding of 0.01 M EDTA solution (1:10, v/v) (see Note 13). 8. Mix the solution thoroughly by carefully shaking the tube (at least 5 times). 9. Centrifugation: 5 min, 240 × g, 4 °C. 10. Discard the supernatant by aspiration and re-suspend the pellet in HBSS (1:10, v/v). 11. Pool both pellets (steps 5 and 10), i.e. the stromal-vascular cell fraction, and transfer the cell suspension in a 50 ml tube. 12. Add erythrocyte-lysis buffer (1:10, v/v) and incubate after mixing the suspension on ice 10 min at longest (see Note 14). 13. Thereafter the tube is filled up to 50 ml with HBSS. 14. Centrifugation: 5 min, 240 × g, 4 °C. 15. Discard the supernatant by aspiration and re-suspend the pellet in basal medium (1:10, v/v). 16. This suspension is filtered without pressure through a 150 μm filter (Image 3). 17. Thereafter flow-through is filtered without pressure through another filter (75 μm) (see Note 15). 18. Centrifuge the flow-through: 5 min, 240 × g, 4 °C (see Note 16).

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Fig. 2 Digested and centrifuged adipose tissue resulting in floating mature adipocytes

Fig. 3 Adipocyte-free pellet before (left) and after erythrocyte lysis (right; the pellet should be white and free of erythrocytes)

19. Discard the supernatant by aspiration and re-suspend the pellet in DMEM/F-12 medium supplemented with 20 % FCS (1:1, v/v) (see Note 17). 20. Inoculate the cells into a 35 mm dish at a density of approx. 50,000 cells/cm2. 3.3 Differentiation of Pre-adipocytes

1. Incubate the cells in a humidified 5 % CO2 atmosphere at 37 °C. 2. Incubate the cells in cultivation medium until reaching confluence.


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Fig. 4 150 μm filter (left) and 75 μm mesh (right)

3. After that the cells are repeatedly washed with 1× PBS to remove non-adhering material and incubated in differentiation medium #1 for another 3 days. 4. Thereafter the cells are washed with 1× PBS to remove nonadhering material and incubated in differentiation medium #2 (see Note 18). 5. The cells are differentiated after at least 7 days. 6. The mature adipocytes can now be cultivated for up to 14 days before becoming apoptotic. 7. In the phase of establishment a lipid staining with oil red is recommended. 3.4 Quality Control of Differentiation by Oil Red O Protocol (Fig. 5)

Oil Red O protocol is an assay performed to detect mature adipocytes in histological visualization of fat cells by staining neutral fat 1. Soak off the culture medium and wash the cells very gently with 1× PBS two times. 2. Discard the supernatant by aspiration and add the fixationsolution. 3. Incubate at RT for 2 h (see Note 19). 4. Discard the fixation solution and wash with 1× PBS two times (see Note 20). 5. Accordingly incubate the cells in 40 % isopropyl alcohol for 5 min. 6. Discard the isopropyl-solution and add the Oil Red O solution.

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Fig. 5 Oil Red O staining of differentiated pre-adipocytes (×400)

7. Incubate at RT for 15 min (see Note 21). 8. Discard the Oil Red O solution and perform a short incubation (~5 s) in isopropyl alcohol. 9. Discard the isopropyl solution and subsequently wash briefly with ddH2O. 10. The wet cells are incubated with the hemalaun solution for 2 min (see Note 22). 11. Afterwards the staining solution is discarded and tap water is added (see Note 23). 3.5 Collecting Secreted Peptides/ Proteins

1. Wash the differentiated pre-adipocytes (mature adipocytes) carefully two times with 1× PBS supplemented with 3 mM CaCl2 (see Note 24). 2. Thereafter add basal medium w/o phenol-red and incubate for 4 h. 3. Wash the cells carefully two times with 1× PBS supplemented with 3 mM CaCl2. 4. Add basal medium w/o phenol-red and incubate the cells for 26 h. 5. Collect the supernatant in 15 ml polypropylene tube and centrifuge for 30 min at 4,900 × g, 4 °C. 6. Transfer the supernatant to another tube, determine the protein concentration and add EDTA solution and protease inhibitors (1×). 7. Concentrate the supernatants with filter concentrator to a protein concentration of 2 μg/μl and store it at −80 °C.


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Fig. 6 Visualization and quality control of secretome samples using 2-dimensional gel electrophoresis. Aliquots (40 μg protein each) of (a) secretome of differentiated, primary adipocytes, (b) total lysate of differentiated primary adipocytes and (c) Fetal Calf Serum (FCS). Samples were separated by 2-dimensional gel electrophoresis according to the 2D-ToGo workflow [9] standard operation procedure established in our group. Accordingly, in the first dimension samples in ReadyPrep Sample Buffer (8 M urea, 3 % CHAPS, 50 mM DTT, 0.2 % (w/v) Bio-Lyte 3/10 ampholytes) were separated by isoelectric focusing (IEF) using pH 3–10 linear ReadyStrip IPGstrips (pH 3–10, 11 cm) performed on a Protean i12™ electrophoresis unit (Bio-Rad) and in the second dimension by SDS-PAGE using Criterion™ TGX Stainfree™ Any KD precast gels (Bio-Rad). After electrophoretic separation spot pattern were acquired and documented without further staining, using a ChemiDoc™ MP (Bio-Rad) equipped with Image Lab Software

3.6 Concentration via Centrifugal Filter Concentrator

Secretome samples frequently are in the range of only a few μg/ml and need a further concentration step to enable comprehensive analysis of the complex secretome. 1. The samples are centrifuged for 45 min at 85,000 × g and 4 °C to get rid of remaining cell debris. 2. The supernatants were transferred to a filter concentrator with small pore size (3 kDa) (see Note 25). 3. Centrifuge at 4,000 × g and 4 °C to a final volume of 100 μl (see Note 26). The secretome sample now is appropriate for comprehensive targeted and non-targeted proteomic profiling. Complexity and sample quality could be evaluated, for example, by 2-DE (Fig. 6), which provides a powerful and reproducible profiling tool.

4

Notes 1. Consider the unique activity of each collagenase-charge: e.g. PZU activity of 1.0 = 1,000 U/mg. When weighing the collagenase wearing a surgical mask is recommended. 2. The CaCl2 is crucial for activation of the collagenase. 3. Do not use this medium for more than a week. 4. TZO must be diluted in DMSO, not in aqueous solvent. 5. The solution should get a violet-blue staining.

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6. The mixture should get a red-violet staining, filtrate the solution with a 150 μm paper filter before use. 7. Only use the solution for 1–2 weeks until precipitation. 8. The buffer has to be freshly prepared, sterile filtered and stored on ice. 9. For rough dissection the use of a forceps and medical scissors is recommended. Therefore, you mince the adipose tissue, e.g. in a weighing pan. 10. Collagenase mixture = tissue/collagenase solution (1:1, v/v). 11. In all of the following centrifugation steps, the use of a swingout rotor is necessary to ensure adequate separation of the fractions. 12. At this point harvested adipocytes can be used for further analysis. On the one hand, it is possible to generate secretome from the floating mature adipocytes and furthermore they can be used for Western blot and qPCR analyses. 13. Do not use a solution of more than 0.5 M of EDTA, because otherwise cells tend to clot. Using serum/FCS is not efficient enough to stop the collagenase reaction. 14. This step is crucial to remove contaminating erythrocytes. It is recommended to monitor the reaction, because ammonium chloride might damage the cells when incubating too long. 15. Both filtering steps are needed to isolate the pre-adipocytes, because using only a single step/mesh might end up with plugging of the mesh and less yield of pre-adipocytes. 16. Never centrifuge for more than 10 min, because the cells are destroyed otherwise. 17. If not planned to work on parallel with the mature adipocytes. 18. This medium was changed every 2–3 days, until full differentiation. 19. This incubation step can be elongated up to 24 h if needed. 20. Wash until the yellowish dye of the washing buffer is extincted. 21. The incubation time depends upon the staining. Better monitor with microscope. 22. For visualizing of the nucleus cells can stained with hemalaun. 23. Use enough water to cover all the cells. Use tap water to ensure presence of HO−-ions which activate the metal–hematoxylin complex. The incubation will probably take longer than 10 min until blue staining. 24. The addition of CaCl2 is crucial to wash away the FCS totally. This enables a subsequent 2D gel analysis. Wash very carefully, because attachment of cells is not very strong.


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25. The use of a 3 kDa filter is of importance to ensure retaining all kinds of proteins in the solution. 26. The retention recovery performance of the filters is ~90 % so that an additional concentration step of the flow-through might be useful. References 1. Misra A, Khurana L (2008) Obesity and the metabolic syndrome in developing countries. J Clin Endocrinol Metab 93:9–30 2. Alberti KG, Eckel RH, Grundy SM et al (2009) Harmonizing the metabolic syndrome: a joint interim statement of the International Diabetes Federation Task Force on Epidemiology and Prevention; National Heart, Lung, and Blood Institute; American Heart Association; World Heart Federation; International Atherosclerosis Society; and International Association for the Study of Obesity. Circulation 120:1640–1645 3. Golay A, Ybarra J (2005) Link between obesity and type 2 diabetes. Best Pract Res Clin Endocrinol Metab 19:649–663 4. Moore JB (2010) Non-alcoholic fatty liver disease: the hepatic consequence of obesity and the metabolic syndrome. Proc Nutr Soc 69:211–220 5. Despres JP, Lemieux I, Bergeron J et al (2008) Abdominal obesity and the metabolic

6.

7.

8.

9.

syndrome: contribution to global cardiometabolic risk. Arterioscler Thromb Vasc Biol 28: 1039–1049 Hauner H, Petruschke T, Russ M et al (1995) Effects of tumour necrosis factor alpha (TNF alpha) on glucose transport and lipid metabolism of newly-differentiated human fat cells in cell culture. Diabetologia 38:764–771 Lehr S, Hartwig S, Sell H (2012) Adipokines: a treasure trove for the discovery of biomarkers for metabolic disorders. Proteomics Clin Appl 6:1–11 Brown KJ, Formolo CA, Seol H et al (2012) Advances in the proteomic investigation of the cell secretome. Expert Rev Proteomics 9: 337–345 Posch A, Franz T, Hartwig S et al (2013) 2D-ToGo workflow: increasing feasibility and reproducibility of 2-dimensional gel electrophoresis. Arch Physiol Biochem 119:108–113

Chapter 5 Methods for Proteomics-Based Analysis of the Human Muscle Secretome Using an In Vitro Exercise Model Mika Scheler, Martin Hrabeˇ de Angelis, Hadi Al-Hasani, Hans-Ulrich Häring, Cora Weigert, and Stefan Lehr Abstract Over the last decade, the skeletal muscle as a secretory organ gained in importance. A growing number of peptides are described which are produced and released by the muscle fibers and work in an autocrine, paracrine, and endocrine fashion. The contraction-induced secretion of these myokines is considered to contribute to the health-promoting effects of exercise. To gain further insights into the molecular processes that occur during contraction an in vitro exercise model, electric pulse stimulation (EPS), was established. Recent publications show that this model is suitable to electro-stimulate human skeletal muscle cells and thus mimic muscle contraction in vitro. Here, we provide a detailed protocol for the proteomics-based analysis of the human muscle secretome, starting with the cultivation of human myotubes and their electric pulse stimulation, ending with sample preparation for targeted and untargeted proteome analysis of the cell culture supernatant. This whole workflow should allow deeper insights into the complex nature of the muscle secretome and the identification of new myokines which might help to understand the crosstalk of the working muscle with different organs and the beneficial effects of exercise. Key words Secretome, Electric pulse stimulation, Myokine, Human myotubes, 2D-DIGE, LC-MS

1

Introduction It is commonly accepted that regular physical activity entails multiple health-promoting effects, plays a key role in the prevention and treatment of metabolic and cardiovascular diseases [1] and reduces systemic inflammation [2]. For decades, it was hypothesized that circulating factors are produced and released from the contracting muscle and enable the communication between the energy-demanding muscle and the energy-supplying organs like adipose tissue and liver during exercise [3]. Interleukin 6 (IL-6) was first discovered as such an “exercise factor” when a higher expression and release of this cytokine was detected in human skeletal muscle after contraction [4] and the putative function of IL-6 as enhancer of endogenous glucose production in exercising


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humans was described [5]. With this cytokine the concept of myokines, peptides produced and released by the muscle fibers which work in an autocrine, paracrine and endocrine fashion arose [6]. Exercise seems to be an important stimulus for the expression and secretion of several myokines including amongst others IL-7, IL-8, Leukemia inhibitory factor (LIF), Myostatin, and Irisin [6]. Although over the last decade many attempts have been made to identify the complex human muscle secretome, the knowledge on myokines and their expression and release especially during contraction is still very limited and incomplete. The global proteomics-based search for exercise factors in human plasma samples is still very challenging due to a concentration range of several orders of magnitude of the different secreted protein species as recently discussed [7]. Proteins and peptides released from muscle fibers are supposed to reach only very low molecular concentrations in the circulation. Certainly, one additional problem is the identification of the muscle cell per se as secretory unit of these circulating exercise factors. Several myokines are not only secreted by the muscle but also by other tissues, e.g. adipose tissue and liver. To overcome these limitations, electric pulse stimulation (EPS) has been applied to human myotubes and the suitability of this model for the identification of myokines was tested. During the last years EPS has been established as in vitro exercise model, which enables the analysis of molecular processes that are occurring during exercise in differentiated skeletal muscle cells. In this model, the in vivo motor nerve activation of the muscle is replaced by electric pulses leading to Ca2+ transients that induce contraction and de novo sarcomere formation in murine C2C12 myotubes and human primary myotubes [8–11]. In human muscle cell culture, the EPS-induced contraction leads to an increased glucose uptake and expression of several proteins involved in oxidative phosphorylation [9, 12]. In a recent paper, we were able to show that EPS of human myotubes leads to an activation of the MAP kinase signaling cascade which results amongst others in an altered myokine expression and that this in vitro exercise model is indeed suitable to analyze the contraction-induced secretome profile of muscle cells [11]. In this study, we compared the transcriptional profile of control cells vs. EPS cells and found 153 significantly upregulated transcripts with fold changes > 1.3 times including 35 genes encoding for secreted proteins. Amongst the top 10 regulated genes with highest fold increases, 7 were supposed to be secreted as proteins. In an immunobead-based assay 12 of 23 analytes were significantly enriched in the EPS supernatant. Using a similar approach, Raschke et al. analyzed conditioned medium of EPS and control cells and identified 48 contraction-induced myokines [13]. These results indicate that the EPS model can be used to study the human muscle secretome by a global proteomics-based approach.

Analysis of the Human Muscle Secretome

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Here, we provide a detailed protocol for the cultivation of human myoblasts, differentiation to myotubes with subsequent electric pulse stimulation and sample preparation of the supernatant for targeted and untargeted proteomics-based analysis.

2

Materials

2.1 General Consumables

1. Pipetboy, pipet and suitable tips. 2. 1.5 mL microcentrifuge tubes. 3. 150 cm2 cell culture dish. 4. 6-well cell culture dish (see Note 1).

2.2

Cell Culture

1. Primary skeletal muscle cells are obtained from percutaneous needle biopsies performed on the lateral portion of quadriceps femoris (vastus lateralis) of human subjects (see Note 2). 2. Cloning medium: 1:1 mixture of α-MEM and Ham’s F-12, 20 % FBS, 2 mM glutamine, 1 % chicken embryo extract, 100 U/mL penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 3. Fusion medium: α-MEM, 2 % FBS, 2 mM glutamine, 100 U/L penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 4. Fusion medium without FBS and phenol red: α-MEM without phenol red, 2 mM glutamine, 100 U/L penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 5. Trypsin solution suitable for cell culture. 6. CO2 incubator (5 % CO2), 37 °C. 7. Phosphate buffered saline (PBS) with and without Mg 2+ and Ca2+.

2.3 Electric Pulse Stimulation

1. Power supply: C-Pace EP Culture Pacer (IonOptix, Dublin, Ireland). 2. Electrodes: C-Dish (IonOptix, Dublin, Ireland).

2.4 Targeted Proteomics

1. Multiplex bead-based immunoassay, e.g. Bio-Plex Pro™ Assay (Bio-Rad, Hercules, CA), Milliplex® Multiplex Assay (Merck Millipore, Darmstadt, Germany). 2. Suitable suspension array system and software for the measurement and data analysis.

2.5 Untargeted Proteomics

1. Falcon tubes (50 mL). 2. Protein concentrators (3 kDa cutoff). 3. Equipment for gel-based or gel-free protein separation technique and MS-based protein analysis [14].

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Methods Cell Culture

1. Thaw, seed, and proliferate human skeletal muscle cells ( see Note 2) in Cloning medium on 150 cm2 plates. Change medium every 2–3 days until cells are 80–90 % confluent. 2. Trypsinate cells and seed at least 20,000 myoblasts per well on 6-well dishes. 3. When cells are 80 % confluent change medium and differentiate cells to multinucleated myotubes (Fig. 1) with Fusion medium for 6–7 days. Change medium every 2–3 days.

3.2 Electric Pulse Stimulation

1. Change medium directly before start of stimulation (see Note 3). 2. Insert electrodes, set culture dish including electrodes into incubator and wire with power supply (Fig. 1; see Note 4). 3. Turn on power supply, adjust the proper program and start stimulation. The intensity and duration of stimulation has to be established for each cell type. We use 14 V, 5 Hz and 2 ms in the basic mode for up to 24 h. The RNA expression of several secreted proteins is increased after less prolonged stimulation, but 24 h are needed to significantly increase glucose consumption and lactate production [11]. Application of higher frequencies (30 Hz) for 4 h leads to an increased phosphorylation of p70S6K1 and expression of MYOD, both depicting high-intensity resistance training leading to increased muscle protein synthesis and muscle hypertrophy [11]. For the proteomics-based investigation of secreted proteins, it is necessary to minimize the contamination with proteins released from damaged cells. The absence of EPS-induced cell damage should be verified by the quantification of cytosolic or mitochondrial localized proteins in the supernatant, e.g. by creatine kinase (CK) activity, also used as plasma marker for muscle fiber damage [15] (see Note 5). The influence of the applied EPS protocol on the cell viability might also be tested using, e.g. the XTT assay. The EPS protocol of 14 V, 5 Hz, 2 ms for 24 h does not decrease the viability of human myotubes or induce the release of CK (Fig. 2a, b; adapted from [11]).

Fig. 1 (continued) myotubes (immunostaining shows MyHC II, a fast-type skeletal muscle myosin in green; nuclei are shown in blue (DAPI staining)). 2. After 6–7 days of differentiation myotubes are electro-stimulated. Therefore, the electrodes dip into the media in the cell culture dish and the whole “sandwich” is put into the incubator. A cable connects the electrode (C-dish) with the power supply (C-Pace EP system) outside (after IonOptix, Dublin, Ireland; detailed brochure: Culture Pacing System). 3. For targeted analysis of the muscle secretome small amounts of media are sufficient whereas untargeted analysis needs larger amounts of media which should be free of FBS and phenol red


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1. Cell culture

Satellite cell isolation

Myoblasts

Myotubes

Power supply

2. EPS

Incubator

3. Proteomics

Normal Fusion media

Targeted immunoassay

Fusion media phenolred FCS

Untargeted approach

Separation (gel-based vs. gel-free)

MS-based analysis for protein identification

Fig. 1 Workflow showing secretome analysis of human myotubes. 1. Skeletal muscle cells are obtained from percutaneous needle biopsies performed on the lateral portion of quadriceps femoris (vastus lateralis) of human subjects. Satellite cells are prepared, myoblasts are cultivated and differentiated to multinucleated

Mika Scheler et al.

Absorbance 450nm (fold change)

a

b 250

1.2

CK activity (U/L)

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1 0.8 0.6 0.4 0.2

200 15 10 5 0

0 con

LDH activity (U/L)

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con

EPS

EPS

SN

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con

EPS

lysate

*

200 15

*

10 5 0 con

EPS

SN

con

EPS

lysate

Fig. 2 EPS does not induce cytotoxicity. Cell viability was measured by XTT cell viability assay (a). Measured absorbance at 450 nm of supernatant is shown as fold change of EPS vs. control supernatant (means ± SEM; n = 5). Creatine kinase (CK; b) and Lactate dehydrogenase (LDH; c) activities were measured in the supernatant (SN, filled bars) and in Triton X-100 lysates of the cells (striped bars). Values are shown in U/L (means ± SEM; n = 5–8; p < 0.05) adapted from [11]

3.3 Collection of Medium for Targeted Secretome Studies

Usually cell culture medium contains FBS to stimulate cell proliferation or in case of serum-reduced media like Fusion media to maintain the viability of the cell. The high abundance of serum proteins in the FBS requires additional enrichment of low-abundant proteins, e.g. using Proteominer technology or metabolic labeling with deuterated amino acids [16, 17]. An efficient and sensitive way to analyze protein signatures in FBS containing supernatant is via commercially available multiplex bead-based immunoassays (see Notes 6 and 7). The following steps are necessary to collect conditioned medium and prepare it for measuring with multiplex bead-based immunoassays. 1. Directly after end of stimulation, take off medium and put on ice until proceeding with the next step (see Note 8). 2. To remove detached cells, spin supernatant with 2,700 × g for 4 min at 4 °C. 3. Transfer supernatant in a new tube and store at −80 °C until used for further analysis.


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4. When using a validated multiplex kit, proceed according to the manufacturer’s instructions (see Notes 8–10). 5. Protein concentrations of the analytes can be calculated with the appropriate optimized standard curves using the appropriate software. 3.4 Collection of Medium for Untargeted Proteomics

An untargeted proteomic profiling approach enables the discovery of novel contraction-induced myokines and helps to identify specific and sensitive biomarkers secreted from the muscle. Untargeted approaches can either be performed by gel-based separation techniques, e.g. 1- and 2D electrophoresis and subsequent staining of the gels or by gel-free separation via HPLC or CE. Proteins can be visualized on gels by fluorescence labeling before electrophoresis (e.g. DIGE) or by staining of the gels after electrophoresis (e.g. silver staining). In all cases, subsequent protein analysis is usually MS-based [14]. In the following part the sample preparation for both separation techniques is described. For LC-MS/MS only small samples sizes are needed whereas for 1- or 2D electrophoresis huge amounts of protein are necessary (see Note 11). Nevertheless, latter application is the method of choice for visualization of the sample composition, its complexity and quality. 1. For untargeted profiling Fusion medium without FBS has to be used during stimulation to prevent the contamination of proteins included in the serum. To decrease any serum carryovers wash cells 2× with PBS containing Ca2+/Mg2+ and 1× with standard PBS (see Note 12). 2. Use 2 mL/well Fusion medium without FBS and phenol red (see Note 13) and start stimulation according to Subheading 3.2. 3. Directly after stimulation end, take off supernatant and spin down detached cells with 800 × g for 5 min at 4 °C to remove any cell debris. 4. Remaining insoluble material is separated by a further centrifugation step (45,000 × g, 5 min, 4 °C). 5. Supernatant can be frozen at −80 °C for further proceeding at this point. 6. Either continue directly with the unfrozen supernatant or thaw the supernatant on ice. Since the native protein concentration is in the lower μg/mL range, the supernatant has to be concentrated by spin concentration devices (cutoff 3 kDa) until a protein concentration of 1–3 mg/mL is reached (see Note 14). 7. Concentrated supernatant is stored at −80 °C until further analysis via gel-based or gel-free protein separation technique and subsequent MS-based protein analysis [14].

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Notes 1. Be sure to use 6-well dishes that fit with the C-dishes (electrodes). The IonOptix handbook proposes 6-well dishes of several companies. To avoid contaminations, heat the electrodes at 100 °C for 3 h. After experiment soak electrodes in demineralized water for several days to neutralize from salts that might have formed on the carbon electrodes and sterilize again. 2. Experiments are performed on the first and second passage of subcultured cells. 3. We use 2 mL medium per well (6-well dish). Be sure that the electrodes dip into the medium and do not contact the bottom of the dish containing the cells. 4. Be careful that the cable is not kinked by the incubator door since it can be damaged then. 5. Lactate dehydrogenase (LDH) is another marker for muscle fiber damage [15]. After 24 h of EP-stimulation, LDH is significantly increased in the total cell lysates of EPS cells and also in the supernatant. These data suggest that elevated levels of cytosolic proteins in the secretome needs to be considered carefully and do not necessarily indicate higher percentage of damaged cells (Fig. 2c; adapted from [11]). 6. For the selection of candidate myokines it might be helpful to do a whole genome transcriptome analysis or quantitative Real-time PCR prior to secretome study to find EPS-regulated proteins. 7. The multiplex assays have several advantages. Only a small sample volume is necessary due to its high sensitivity. Additionally, it is easy to handle, enables an exact quantification in a concerted system. The biggest advantage compared to commercial available enzyme-linked immunosorbent assay (ELISA) is the measurement of several proteins simultaneously in one assay. 8. During immunoassay antibodies might cross-react with proteins of FBS, hence a sample of unconditioned medium (reference medium) is urgently needed for multiplex measurement. Spin down the reference medium according to methods in step 2 in Subheading 3.3 and store supernatant at −80 °C until used for further analysis. 9. When mixing individual singleplexes check x-Plex Bead Regions and avoid using 2 singleplexes with similar x-Plex Bead Regions in one assay. 10. For the conditioned medium of human myotubes, neat samples were used (50 μL sample/well). In our case, all measured ana-


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lytes were in the assay working range. If you are above the upper limit of quantification make sure to dilute the samples properly. 11. For comprehensive analysis of the secretome via 2DE, a protein concentration up to mg/mL range is required. In the special case of DIGE technology at least 150 μg protein is essential. Thus, huge amounts of medium are necessary. 12. To get rid of the complex protein mixture included in FBS, cells are washed 2 times with PBS containing Ca2+/Mg2+. This is used to wash adherent cell cultures when one merely wishes to wash and have the cells remain adherent because adhesion proteins require divalent cations to function. The third washing step is done with PBS without Ca2+/Mg2+ to decrease salt concentration in the medium. 13. For untargeted proteomics the medium should be free of phenol red because it might interfere with protein assays performed to determine protein concentration for subsequent gels. 14. Since the native protein concentration is very low, it is essential to concentrate the supernatant to achieve a working concentration for subsequent gels. It might be necessary to transfer the supernatant during concentration process in an even smaller protein concentrator tube to gain smaller volumes and the required protein concentration. Be aware that with every change of the device material gets lost. To minimize the loss, rinse the membranes of the larger device with flow through and add it to the eluent of the smaller device. Additionally, take care of protein precipitations that might form.

Acknowledgements This study was supported in part by a grant from the Leibniz Gemeinschaft (SAW-FBN-2013-3) to C. Weigert and by a grant from the German Federal Ministry of Education and Research (BMBF) to the German Center for Diabetes research (DZD e.V.; No. 01GI0925). References 1. Pedersen BK (2009) The diseasome of physical inactivity—and the role of myokines in muscle— fat cross talk. J Physiol 587:5559–5568 2. Gleeson M (2007) Immune function in sport and exercise. J Appl Physiol (1985) 103:693–699 3. Goldstein MS (1961) Humoral nature of the hypoglycemic factor of muscular work. Diabetes 10:232–234

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17.

human skeletal muscle cells as an in vitro model of exercise. PLoS One 7:e33203 Raschke S, Eckardt K, Bjorklund Holven K et al (2013) Identification and validation of novel contraction-regulated myokines released from primary human skeletal muscle cells. PLoS One 8:e62008 Wohlbrand L, Trautwein K, Rabus R (2013) Proteomic tools for environmental microbiology—a roadmap from sample preparation to protein identification and quantification. Proteomics 13:2700–2730 Fernandez-Gonzalo R, Lundberg TR, AlvarezAlvarez L et al (2014) Muscle damage responses and adaptations to eccentricoverload resistance exercise in men and women. Eur J Appl Physiol 114(5): 1075–1084 Frobel J, Hartwig S, Passlack W et al (2010) ProteoMiner and SELDI-TOF-MS: a robust and highly reproducible combination for biomarker discovery from whole blood serum. Arch Physiol Biochem 116:174–180 Eichelbaum K, Winter M, Berriel Diaz M et al (2012) Selective enrichment of newly synthesized proteins for quantitative secretome analysis. Nat Biotechnol 30:984–990

Chapter 6 Urinary Pellet Sample Preparation for Shotgun Proteomic Analysis of Microbial Infection and Host–Pathogen Interactions Yanbao Yu and Rembert Pieper Abstract Urine is one of the most important biofluids in clinical proteomics, and in the past decades many potential disease biomarkers have been identified using mass spectrometry-based proteomics. Current studies mainly perform analyses of the urine supernatant devoid of cells and cell debris, and the pellet (or sediment) fraction is discarded. However, the pellet fraction is biologically of interest. It may contain whole human cells shed into the urine from anatomically proximal tissues and organs (e.g., kidney, prostate, bladder, urothelium, and genitals), disintegrated cells and cell aggregates derived from such tissues, viruses and microbial organisms which colonize or infect the urogenital tract. Knowledge of the function, abundance, and tissue of origin of such proteins can explain a pathological process, identify a microbe as the cause of urinary tract infection, and measure the human immune response to the infection-associated pathogen(s). Successful detection of microbial species in the urinary pellet via proteomics can serve as a clinical diagnostic alternative to traditional cell culture-based laboratory tests. Filter-aided sample preparation (FASP) has been widely used in shotgun proteomics. The methodology presented here implements an effective lysis of cells present in urinary pellets, solubilizes the majority of the proteins derived from microbial and human cells, and generates enzymatic digestion-compatible protein mixtures using FASP followed by optimized desalting procedures to provide a peptide fraction for sensitive and comprehensive LC-MS/MS analysis. A highly parallel sample preparation method in 96-well plates to allow scaling up such experiments is discussed as well. Separating peptides by nano-LC in one dimension followed by online MS/MS analysis on a Q-Exactive mass spectrometer, we have shown that more than 1,000 distinct microbial proteins and 1,000 distinct human proteins can be identified from a single experiment. Key words Urine proteomics, Urinary pellet, Microbial infection, Host–pathogen interaction, Filter-aided sample preparation (FASP), 96FASP

Abbreviations FASP HCD LC

Filter-aided sample preparation Higher energy collision disassociation Liquid chromatography


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MS MWCO TIC UTI

1

Mass spectrometry Molecular weight cut off Total ion chromatogram Urinary tract infection

Introduction Urine is a sample source of high importance for biomarker discovery because it is easily available and collected non-invasively in large quantities [1, 2]. The identity and quantity of proteins excreted into urine may reflect pathological conditions that can be traced to different organs in the body, particularly the kidneys, prostate, and urogenital tract [3]. Currently, most urine proteomic studies focus on the analysis of the urinary supernatant, that is the soluble fraction of the collected urine sample following centrifugation at 1,500–5,000 × g for 5–15 min [4, 5]. The resulting urinary pellet is frequently discarded. However, the urinary pellets, especially those from patients with urinary tract infection (UTI), which is one of the most common conditions that lead to hospital visits [6], contain not only pathogenic microbes, in most cases bacteria that colonize the urinary tract of the patient, but also host proteins associated with the inflammatory process following colonization with the microbial pathogen. Inflammation may include activities such as recognition of pathogen molecular patterns, cytokine release, leukocyte recruitment, lymphocyte recruitment, complement activation, immunoglobulin secretion, fibrin deposition, release of iron-sequestering proteins and direct microbial killing via enzymatic activities and permeabilization or disintegration of bacterial membranes [6–8]. The presence and relative quantity of such proteins serve as a diagnostic indicator of infection and inflammation [7]. Non-pathogenic bacteria not inducing inflammatory responses may also be identified from urinary pellets [9]. Our laboratory reported the first metaproteomic analysis of urinary pellets derived from patients diagnosed with either asymptomatic bacteriuria or UTI, and identified the microbial causes of bacteriuria [9]. Most recently, our laboratory developed a highthroughput urinary sample preparation approach, 96FASP (96well filter-aided sample preparation), for quantitative shotgun proteomic analysis [10]. This method promises to be the prototype of an economical method for the diagnosis of urinary tract infections and inflammation in the future. This article describes an extensively used, robust step-by-step procedure pertaining to the preparation of urinary pellet samples using the FASP approach. The optional 96FASP method that is adapted to process multiple samples simultaneously is described as well.

Urinary Pellet Proteomics

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Materials

2.1 Cell Lysis and FASP

1. Sartorius Vivacon 500 filter device (30,000 MWCO). 2. MultiScreen Ultracel-10 Filter Plate 10 kDa (Merck-Millipore, USA). 3. Bench-top centrifuge (for example, Eppendorf 5415R or equivalent). 4. (Optional) Plate centrifuge (for example, Eppendorf 5810R or equivalent). 5. Misonix Sonicator 3000 Ultrasonic Cell Disruptor. 6. Pre-casted SDS PAGE gel (for example, NUPAGE 4–12 %). 7. SpeedVac (for example, Labconco Refrigated CentriVap Concentrator, or equivalent). 8. TMN buffer: 40 mM Tris–HCl, pH 8.1, 5 mM MgCl2 and 100 mM NaCl. 9. Lysostaphin solution: 10 mg/ml in water (AMBI Products; from Staphylococcus simulans). 10. Mutanolysin solution: 2 mg/ml in water (Sigma-Aldrich; from Streptococcus globisporus). 11. NaOH solution: 100 mM in water. 12. UA buffer: 8 M urea in 50 mM Tris–HCl, pH 8.1. UA buffer should be prepared freshly each day. 13. USED lysis buffer: 8 M Urea, 1 % SDS, 5 mM Na-EDTA, 50 mM DTT. USED buffer should be prepared freshly each day. 14. IAA solution: 0.05 M iodoacetamide in 50 mM Tris–HCl, pH 8.1. IAA solution should be prepared freshly each day. 15. ABC buffer: 50 mM ammonium bicarbonate in water. 16. Trypsin solution: trypsin (Promega sequencing grade); stock concentration 0.1 μg/μl and stored at −80 °C.

2.2 StageTip Desalting

1. Empore C18 Extraction disks (3 M, catalog number: 2215). 2. Activation buffer: 100 % methanol. 3. Wash and equilibration buffer: 0.5 % acetic acid in H2O. 4. Elution solution-I: 0.5 % acetic acid, 60 % acetonitrile, and 40 % H2O. 5. Elution solution-II: 0.5 % acetic acid, 80 % acetonitrile, and 20 % H2O.

2.3

LC-MS/MS

1. XCalibur software (version 2.2, Thermo Scientific). 2. Proteome Discoverer (version 1.4, Thermo Scientific). 3. HPLC solvent A: 0.1 % formic acid in water. 4. HPLC solvent B: 0.1 % formic acid in acetonitrile.

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5. Trap column: C18 PepMap100, 300 μm × 5 mm, 5 μm, 100 Å (Thermo Scientific, USA). 6. Analytical column: PicoFrit, 75 μm × 10 cm, 5 μm BetaBasic C18, 150 Å (New Objective, USA). 7. Ultimate 3000 nano-LC system coupled to Q-Exactive mass spectrometer (Thermo Scientific, USA).

3

Methods An overview of the protein sample preparation for urinary pellets is provided in Fig. 1, as explained in detail in the following procedures. The schematic also shows the downstream applications (e.g., LC-MS/MS and database search) for urinary proteome analysis.

3.1 Lysis of Urinary Pellets and Collection of Lysates

1. Take urine samples (see Note 1), centrifuge at 3,000 × g for 15 min at 4 °C, discard the urine supernatant and recover the pellet, retaining ~1.0 ml of residual urine supernatant to avoid disturbing the pellet. 2. Add ~10 ml ice-cold phosphate-buffered saline PBS, gently shake the tube and centrifuge for another 15 min at 3,000 × g. Discard the supernatant and store the wet pellet in −80 °C or process immediately. 3. Add USED buffer (see Note 2) consistent with an approximate volume ratio of 4:1 (buffer volume/urinary pellet volume) to lysis cells and solubilize the contents in the urinary pellet. If the urinary pellet volume is very small, a minimum of 100 μl USED buffer volume can be added. 4. Vortex vigorously for 10 s a few times to resuspend the pellet; pipette up and down to resuspend the pellet if the vortexing step was not sufficient to homogenize the pellet. Incubate for 30 min at room temperature and vortex or flick tube occasionally. 5. Using Misonex Sonicator (with the water bath attached, not the probe), put the urinary lysate into an ice/water-filled water bath. Set the program on amplitude 9 for nine 45-s cycles with 45-s intermittent cooling of the urinary lysate sample(s). 6. Let the urinary lysate sample(s) sit for approximately five more minutes. Spin urinary lysate sample(s) in a bench-top centrifuge at maximum speed (14,000 × g) for 10 min, and transfer lysate supernatant to a new 1.7 ml microtube (see Note 3). Freeze the urinary lysate at −80 °C if necessary.

3.2 FASP Processing Urinary Pellet Samples for Shotgun Proteomics

1. Before processing pellet lysate using FASP, flush the filters with 200 μl NaOH (100 mM) and 200 μl UA buffer once separately. 2. Aliquot a volume equivalent to 10–100 μg total protein quantity into Vivacon 500 filter device, and mix with 200 μl UA buffer.


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Fig. 1 An overview of the urinary pellet sample preparation for shotgun proteomics. The procedures are explained in detail in this chapter. Briefly, the urine samples are first spin down to collect pellet fraction. The pellets are then lysed and digested following FASP protocol. A protocol of 96-well format FASP is also illustrated. Afterwards, the protein digests are cleaned using StageTip and analyzed by nanoLC-MS/MS. The host and pathogen protein identifications can be obtained by searching a metaproteome database which includes human proteins as well as the proteins of common pathogens related to your study (in this case, the urinary tract infection)

3. Spin at 14,000 × g for 10–30 min. 4. Add 200 μl UA buffer and repeat the spin at 14,000 × g to concentrate until the volume in the filter unit is reduced to ~10 μl. 5. Add 100 μl of the IAA solution (final concentration 50 mM) and mix by vortexing filter unit for 1 min. Incubate without mixing for 20 min in the dark at ambient temperature. 6. Spin at 14,000 × g for 10–30 min. Discard flow-through from collection tube.

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7. Add 200 μl of UA buffer and spin at 14,000 × g for 10–30 min. The final sample volume in the filter unit should be 20 μl or less. 8. Add 200 μl of ABC buffer and spin at 14,000 × g for 10–15 min. Add 200 μl of ABC buffer and repeat spin step once more. 9. Add 100 μl of ABC buffer to filter unit and trypsin so that the final protein:trypsin ratio is 100:1. Mix by vortexing for 1 min. Transfer the filter to a new filter collection tube. 10. Incubate at 37 °C overnight in a temperature-controlled incubator (12–16 h). 11. On the next day, add 200 μl of ABC buffer, spin the filter unit at 14,000 × g for 10–15 min and collect the filtrate (with the peptide mixture) into a maximum recovery tube. 12. Add 100 μl of ABC buffer to filter unit and trypsin so that the final protein:trypsin ratio is 100:1, and re-incubate the tube at 37 °C for another 4–6 h. 13. Add 200 μl of ABC buffer, spin the filter unit after this second digestion step at 14,000 × g for 10–15 min and collect the second filtrate (with the peptide mixture) into the same maximum recovery tube. 14. Add another 200 μl of ABC buffer, spin the filter unit after the wash step at 14,000 × g for 10–15 min and collect the wash (with residual peptides) into the same maximum recovery tube. The final volume in the collection tube is ~600 μl. 15. Dry the peptide solution using a Speed-Vac (this may take 2–3 h). The tryptic peptide mixture is ready now for desalting with the StageTip method. 3.3 (Optional) 96-Well Filter Plate Processing Urinary Pellet Samples for Shotgun Proteomics

1. Instead of using individual Vivacon filters to process urinary pellet samples separately, one can use 96-well filter plate to process multiple samples simultaneously. This could be a good option when tens or hundreds of samples have to be analyzed [10]. 2. All the following procedures are the same as described above for single filter processing; there are a couple of changes: (a) Each spin takes place on plate centrifuged at 2,600 × g for 45–90 min. (b) The collection plate should be polypropylene-based (e.g., to reduce possible non-specific bindings). (c) The lid of the 96-well filter plate cannot seal the plate very well; to avoid sample drying out after overnight incubation, parafilm could be used to warp and tightly seal the lid. In the meantime, at least 100 μl ABC buffer should be added into each well for overnight digestion.


3.4 Peptide Desalting Using StageTip Protocol

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1. This method is adapted from a published protocol [11]; several changes have been made to optimally fit the preparation of urinary pellet samples (see Note 4). 2. Prepare StageTip by punching out small discs (1–3 layers depending on the sample amount) of C18 Empore filter using a 22 G flat-tipped syringe and ejecting the discs into P200 pipette tips. Ensure that the disc is securely wedged in the bottom of the tip. 3. Activate a tip by forcing 200 μl methanol through the tip. Use this step to check if the StageTip is leaky or overtight. 4. Force 200 μl Elution Solution-II through the tip. 5. Equilibrate tip by forcing 200 μl Wash and equilibration buffer through the tip. The tip is now ready for sample loading. 6. Resuspend the dried peptides into 100 μl of Wash and equilibration buffer, and vortex for 10 min. 7. Load the 100 μl peptide solution—made in step [5] above— by forcing them through the C18-StageTip. Do not discard the flow-through, but collect it into the original tube. 8. Reload the peptide solution onto the tip. This step may be repeated two or three times. 9. Wash the tip with 200 μl Wash buffer, and repeat this step one or two times. The flow-through during this step can be discarded. 10. Elute the peptides with 200 μl Elution Solution-I (once) and 200 μl Elution Solution-II (twice). Collect all of the eluates (~600 μl) into one maximum recovery microtube. 11. Dry the peptide eluates in the Speed-Vac (this may take 1–2 h). 12. Store then at −80 °C, or resuspend it with LC solvent A for immediate LC-MS/MS analysis.

3.5 Nano-LC-MS/MS and Computational Analysis

1. Resuspend the dried peptide samples into 20–50 μl LC solvent A, centrifuge at maximal speed for 3–5 min, transfer to sample vials and load to HPLC autosampler. 2. The LC-MS/MS analysis is operated by an Ultimate 3000 nano LC system and Q Exactive mass spectrometer. Around 2–5 μl of the samples were first loaded onto a trap column at high flow rate 5 μl/min, and then separated on a PicoFrit analytical column at a nano flow rate of 300 nl/min. For a 2-h LC-MS run, a linear gradient was applied from 100 % solvent A to 35 % solvent B over 100 min, followed by a steeper gradient to 80 % solvent B over 15 min. The column was re-equilibrated with solvent A for 5 min. Eluting peptides were acquired in data-depended mode using XCalibur software and top10 method as described before [10].

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3. The acquired raw files are then processed using the Proteome Discoverer software. The protein database involved in this study contains UniProt human protein sequences and common urinary tract pathogens [12]. They can be downloaded from UniProt website (http://www.uniprot.org/). MS search parameters are similar to published previously [10]. 4. Figure 2 shows a typical nanoLC-MS/MS analysis of the urinary pellet samples. The base peak of the TIC shows the eluted peptides from HPLC column are acquired by mass spectrometer in 120 min. Shown in the upper panel is a representative full MS scan collected at retention time 75.28 min. The mass of each peak (peptide) are accurately measured by the Orbitrapbased mass analyzer at high resolution (e.g., 70,000), and then sequenced in the HCD-based collision cell. The resulting

Fig. 2 A typical LC-MS analysis of the urinary pellet sample. The full mass (MS) and fragmentation (MS/MS) are recorded by high resolution and high accuracy mass spectrometer. A representative MS scan at a given time (for example, t = 75.3 min, as indicated by the red bar and arrows in the lower panel) with mass and charge state information (for example, m/z = 668.3510, z = 2) is shown in the middle panel. Most of the ions in this spectrum will be isolated and fragmentized afterwards. Then, the bioinformatics tool (for example, Sequest algorithm) will assign the most confident amino acid sequence to each peak based on the sophisticated scoring system. Detailed explanations of the figure are provided in this chapter


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fragments are measured again by the Orbitrap analyzer with high resolution (e.g., 17,500). With high quality MS and MS/ MS data (mass errors are shown for each peak in the figure), database search engine (e.g., Sequest) will assign each peak a unique amino acid sequence and its corresponding protein, as indicated by triangles (red, E.coli proteins; black, human proteins) in the MS scan. The peak m/z = 846.0721 is magnified in the right window. The cysteine in protein TRFL has a carbamidomethyl modification. The gray diamond ( ) shows a keratin peptide (VDALMDEINFMK, ∆ = −2.74 ppm; [K2C5_HUMAN]). In this single LCMS analysis, 2,198 proteins, including human (1,139) and bacterial proteins (1,059), were successfully identified.

4

Notes 1. Usually 5–50 ml of urine is collected from human subjects. The preferred approach is to process the urine sample immediately after collection by centrifugation (see step 2). It is possible to store the urine at 4 °C for up to 6 h before centrifugation. Finally, the entire urine sample may be stored at −80 °C at the clinical site, shipped to the site of proteomic analysis and thawed prior to centrifugal separation of urinary supernatant and urinary pellet. The freeze–thaw step may alter the composition of the urinary pellet. For a given project with multiple samples, a distinct urine storage and processing method should be selected. 2. In the case that gram-positive bacteria with thick cell walls, such as Streptococcus pneumoniae and Staphylococcus aureus, are suspected to be present in urine samples, resuspend the urinary pellet samples in a volume of TMN buffer to have an approximate volume ratio of 1:10, usually less than 200 μl. Pipette the suspension up and down a few times in a 1.5 ml tube. Add lysostaphin and mutanolysin to a final concentration of 20 μg/ml. Mix gently to homogenize the enzymes and suspended cells. Incubate the sample in the 37 °C shaker-incubator. Take out the tube every 30 min and check if a pellet collects at the bottom. If so, briefly vortex every 30 min. Complete digestion after a 3 h incubation. After pre-treatment with lysostaphin and mutanolysin, add up to 600 μl USED buffer into the lysate. 3. To estimate the protein concentration, take 10 μl aliquot of the supernatant to another new microtube, mix with SDS loading buffer and run it in a SDS-PAGE gel. Load 2 and 5 μg BSA standards in the same gel. Coomassie Blue (CB)-G250 stain the gel followed by its destaining with standard procedures [13]. From the overall CB-G250 staining intensity of urinary pellet lysate bands, estimate the total protein amount

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in the lane based on BSA staining intensities. The protein concentration could also be measured by tryptophan fluorescence as reported before [14]. 4. Instead of using syringe to manually push solvents through the StageTips, one can use pipette tip adaptors (commercially available from The Nest Group, MA) which fit the 1.5- or 2.0mL microtubes well. This way, all the processing steps with syringe can now be done using a bench-top centrifuge [15].

Acknowledgments This work was supported in part by Grant NIH-1R01GM103598 (National Institute of General Medical Sciences). References 1. Decramer S, de Peredo AG, Breuil B, Mischak H, Monsarrat B, Bascands J-L, Schanstra JP (2008) Urine in clinical proteomics. Mol Cell Proteomics 7:1850–1862 2. Wood SL, Knowles MA, Thompson D, Selby PJ, Banks RE (2013) Proteomic studies of urinary biomarkers for prostate, bladder and kidney cancers. Nat Rev Urol 10:206–218 3. Barratt J, Topham P (2007) Urine proteomics: the present and future of measuring urinary protein components in disease. Can Med Assoc J 177:361–368 4. Adachi J, Kumar C, Zhang Y, Olsen J, Mann M (2006) The human urinary proteome contains more than 1500 proteins, including a large proportion of membrane proteins. Genome Biol 7:R80 5. Rodríguez-Suárez E, Siwy J, Zürbig P, Mischak H (2014) Urine as a source for clinical proteome analysis: from discovery to clinical application. Biochim Biophys Acta 1844:884–898 6. Nielubowicz GR, Mobley HLT (2010) Hostpathogen interactions in urinary tract infection. Nat Rev Urol 7:430–441 7. Pieper R, Suh M, Fouts D, Nelson K (2012) Metaproteomic method for diagnosis of bacteriuria, urogenital tract and kidney infections from urinary pellet samples. US Patent App. 13/728,106 8. Weichhart T, Haidinger M, Hörl WH, Säemann MD (2008) Current concepts of molecular defence mechanisms operative during urinary tract infection. Eur J Clin Invest 38:29–38

9. Fouts D, Pieper R, Szpakowski S, Pohl H, Knoblach S, Suh M-J, Huang S-T, Ljungberg I, Sprague B, Lucas S, Torralba M, Nelson K, Groah S (2012) Integrated next-generation sequencing of 16S rDNA and metaproteomics differentiate the healthy urine microbiome from asymptomatic bacteriuria in neuropathic bladder associated with spinal cord injury. J Transl Med 10:174 10. Yu Y, Suh M-J, Sikorski P, Kwon K, Nelson KE, Pieper R (2014) Urine sample preparation in 96-well filter plates for quantitative clinical proteomics. Anal Chem 86:5470–5477 11. Rappsilber J, Mann M, Ishihama Y (2007) Protocol for micro-purification, enrichment, pre-fractionation and storage of peptides for proteomics using StageTips. Nat Protoc 2:1896–1906 12. Foxman B (2010) The epidemiology of urinary tract infection. Nat Rev Urol 7:653–660 13. Shevchenko A, Tomas H, Havlis J, Olsen JV, Mann M (2007) In-gel digestion for mass spectrometric characterization of proteins and proteomes. Nat Protoc 1:2856–2860 14. Wiśniewski JR, Duś K, Mann M (2013) Proteomic workflow for analysis of archival formalin-fixed and paraffin-embedded clinical samples to a depth of 10 000 proteins. Proteomics Clin Appl 7:225–233 15. Yu Y, Smith M, Pieper R (2014) A spinnable and automatable StageTip for high throughput peptide desalting and proteomics. Protoc Exchange. doi:10.1038/protex.2014.033

Chapter 7 A Protocol for the Parallel Isolation of Intact Mitochondria from Rat Liver, Kidney, Heart, and Brain Sabine Schulz, Josef Lichtmannegger, Sabine Schmitt, Christin Leitzinger, Carola Eberhagen, Claudia Einer, Julian Kerth, Michaela Aichler, and Hans Zischka Abstract Mitochondria are key organelles for cellular energy production and cell death decisions. Consequently, a plethora of conditions which are toxic to cells are known to directly attack these organelles. However, mitochondria originating from different tissues differ in their sensitivity to toxic insults. Thus, in order to predict the potential organ-specific toxicity of a given drug or pathological condition at the mitochondrial level, test settings are needed that directly compare the responses and vulnerabilities of mitochondria from different organs. As a prerequisite for such test strategies, we provide here a robust, prompt, and easy-tofollow step-by-step protocol to simultaneously isolate functional and intact mitochondria from rat liver, kidney, heart, and brain. This isolation procedure ensures mitochondrial preparations of comparable purity and reproducible quantities which can be subsequently analyzed for organ-specific mitochondrial toxicity. Key words Mitochondria, Liver, Heart, Brain, Kidney

1

Introduction The prime role of mitochondria in cell metabolism reflects the fact that these organelles are crucial in controlling the cells’ fate, i.e. survival or death, most importantly by balancing the cellular energy demands. However, metabolic differences and metabolite preferences exist in the different healthy tissues of our body [1]. Whereas brain tissue relies on glucose as the major metabolite, liver, especially in the postresorption phase, relies on fatty acids [1]. Consistent with these metabolic preferences, marked differences in the molecular composition of the respective mitochondrial populations are known [2–4]. Thus, mitochondria delicately adapt at the molecular level to the metabolic state of their tissue origin.


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Consequently, upon exposure to toxic insults that target mitochondria, these organelles may respond differently and in an organ-specific fashion. For example, succinate is an important substrate for mitochondrial ROS production in brain, heart, kidney, and skeletal muscle mitochondria whereas fatty acids generate significant quantities of oxidants in kidney and liver mitochondria [5]. Moreover, liver and brain mitochondria dramatically differ in their response toward calcium challenges [6] and, importantly, were shown to significantly differ in their sensitivity toward elicit of the mitochondrial permeability transition, a cell death causing event [7]. Thus, severe tissue-dependent differences in the mitochondrial sensitivities toward detrimental impacts exist. In order to investigate such organ-specific mitochondrial differences, test strategies are needed that minimize inter-experimental variability. Preferably, mitochondria to be compared should be isolated in parallel on the same day, under appropriate conditions each, and from the same animal in order to subsequently and simultaneously analyze them for their individual susceptibilities. Here we provide a protocol to isolate mitochondria retaining their metabolic functionality, from the same animal simultaneously and from at least four different rat tissues, namely liver, kidney, heart, and brain. This protocol is a combination of classical isolation procedures for mitochondria from individual sources based on differential centrifugation, introduced by Palade’s group [8] but containing several adaptations according to published methods [9]. As these “crude” mitochondrial-enriched fractions still contain other relevant cellular compartments, particularly peroxisomes and lysosomes, they are subjected to further Percoll™ gradient centrifugation resulting in significant purification [10]. Such purified mitochondria are comparably homogenous but retain their inherent structural peculiarities depending on their tissue origin (Figs. 1 and 2). Importantly, these isolated mitochondria present with intact membrane structures as they do not only efficiently build up an inner transmembrane potential upon substrate addition (e.g. succinate), but also retain this membrane potential for at least 60 min, highly indicative of preserved functional integrity (Fig. 3). Thus, the mitochondrial populations isolated from the same animal appear similar in sample purity and functional integrity, being therefore highly suitable for subsequent differential sensitivity testing. In total, the isolation procedure takes about 2.5 h (see Note 1) and the tissue-dependent mitochondrial average yield is given in Table 1. This protocol may be adapted to isolations of mitochondria from other tissues, e.g. pancreas or spleen for comparative purposes (see Note 2).

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Fig. 1 Electron micrographs of liver, kidney, heart, and brain mitochondria in situ (Scale bars equal 2 μm for 1,600× magnification, and 500 nm for 10,000× and 20,000× magnifications, respectively). Hepatocyte mitochondria appear round or elongated in shape, with an electron-dense matrix, mostly tubular cristae and dense mitochondrial granules. Kidney tubule cell mitochondria contain densely packed elongated mitochondria oriented in right angles to the base of the cell. Myocardial muscle cell mitochondria are round or elongated in shape comprising numerous, closely arranged cristae. Myelinated axon mitochondria appear with lamellar and tubular cristae

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Fig. 2 Electron micrographs of liver, kidney, heart, and brain mitochondria isolated in parallel by the herein presented protocol from the same animal (scale bars equal 5 μm for 5,000×, 1 μm for 20,000×, and 500 nm for 50,000× magnifications). Homogeneous mitochondrial populations are obtained. However, as observed in situ, depending on their tissue origin isolated mitochondria appear with significant structural differences

2

Materials

2.1 Isolation Procedure Components

1. Isofluran anesthesia facility (Sigma Delta Vaporizer, Penlon, Abingdon, UK) and Isofluran (IsoFlo®, Ecuphar, Greifswald, Germany). 2. Teflon-glass homogenizer (B. Braun Biotech “Homogenisator Potter S”, Melsungen, Germany) with manual hub and compatible pestles for 30 and 15 mL volumes.

Fig. 3 A time stable mitochondrial membrane potential in freshly isolated mitochondria confirms the functional integrity of their inner membranes. Assessment of the mitochondrial membrane potential was done using the fluorescent dye Rhodamine 123 according to published protocols [13]. (a) Due to accumulation within mitochondria and fluorescence quenching of the positively charged dye upon an intact, negative inside membrane potential, a low fluorescence is indicative for an existing membrane potential. Upon addition of the protonophor FCCP as internal control, dissipation of the mitochondrial membrane potential occurs, which can be followed by a steep increase in fluorescence. (b) As a semi-quantitative measure for the time stable mitochondrial integrity the fluorescence ratio before and immediately after FCCP addition is calculated using the time points t = 0 (first enlarged light grey diamond) and t = 60 (second enlarged grey diamond) minutes

Table 1 Average tissue-dependent yield of mitochondria given as mg mitochondrial protein per g tissue wet weight [14] Tissue

Mitochondria [mg/g w.w.]

Animals

Liver

17.7 ± 5.1

11

Kidney

3.9 ± 1.5

11

Heart

1.2 ± 0.6

11

Brain

0.4 ± 0.3

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3. Rough glass-in-glass homogenizer 5 mL volume (Homogenisator LAT, Reiss Laborbedarf e.K., MainzMombach, Germany). 4. Petri dishes, 60 mm diameter (Nunclon®, Sigma-Aldrich, Taufkirchen, Germany). 5. Blade, 39 mm cutting width (NeoLab, Heidelberg, Germany). 6. Centrifugation tubes: 50 mL (Polypropylene, Beckman Coulter, Krefeld, Germany) and 15 mL (transparent, preferably glass, Corex®, DuPont Instruments, Neu-Isenburg, Germany). 7. Percoll™ solution density 1.130 mg/mL (GE Healthcare, Munich, Germany). 8. Prepare all solutions using ultrapure water (Prepared by purifying deionized water to attain a resistivity of 18.2 MΩ-cm at 25 °C). 2.2 Isolation Procedure: Buffers and Solutions

1. 0.9 % NaCl: Weigh 0.9 g NaCl into a 100 mL graduated flask, add water up to 100 mL and let stir at room temperature until NaCl is solved. 2. Isolation buffer with BSA: 0.3 M sucrose, 5 mM TES, 0.2 mM EGTA, 0.1 % BSA (w/v), pH 7.2. Weigh 102.8 g sucrose, 1.146 g TES, 76 mg EGTA and 1 g BSA (fatty acid free) and transfer to a 1 L glass beaker. Add ultrapure water to a volume of ~800 mL and let stir for 30 min at RT to solve all chemicals (see Note 3). Adjust the pH diligently to 7.2 with 5 M KOH at RT (see Note 4). Transfer to a 1 L graduated flask and add water up to 1 L and mix thoroughly. Store isolation buffer with BSA at 4 °C and use within 1 week. 3. Isolation buffer without BSA: 0.3 M sucrose, 5 mM TES, 0.2 mM EGTA, pH 7.2. Weigh 102.8 g sucrose, 1.146 g TES, and 76 mg EGTA and transfer to a 1 L glass beaker. Add ultrapure water to a volume of ~800 mL and let stir for 30 min at RT to solve all chemicals (see Note 3). Adjust the pH diligently to 7.2 with 5 M KOH at RT (see Note 4). Transfer to a 1 L graduated flask and add water up to 1 L and mix thoroughly. Store isolation buffer without BSA at 4 °C and use within 1 week. 4. Isolation buffer for Percoll™ gradients (IPP buffer): 0.3 M sucrose, 10 mM TES, 0.2 mM EGTA, 0.1 % BSA (w/v), pH 6.9. Weigh 51.4 g sucrose, 1.146 g TES, 38 mg EGTA and 0.5 g BSA (fatty acid free) and transfer to a 1 L glass beaker. Add ultrapure water to a volume of ~400 mL and let stir for 30 min at RT to solve all chemicals (see Note 3). Adjust the pH diligently to 6.9 with 5 M KOH at RT (see Note 4). Transfer to a 500 mL graduated flask and add water up to 500 mL and mix thoroughly. Store IPP buffer at 4 °C and use within 1 week.


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5. Prepare gradient solutions A–C as follows: Gradient solution A: supplement 60 mL of IPP buffer with 1.43 g sucrose. Gradient solution B: supplement 60 mL of IPP buffer with 2.65 g sucrose. Gradient solution C: supplement 60 mL of IPP buffer with 9.24 g sucrose. Gradient solutions A–C can be stored at 4 °C for 1 week.

3

Methods

3.1 Preparation of Percoll Gradients

1. Prepare gradient layer solutions with 18, 30, and 60 % Percoll as follows (see Note 5): 18 % gradient layer solution: supplement 24.6 mL of gradient solution A with 5.4 mL Percoll. 30 % gradient layer solution: supplement 21 mL of gradient solution B with 9 mL Percoll. 60 % gradient layer solution: supplement 12 mL of gradient solution C with 18 mL Percoll. 2. Using a Peleus ball (see Note 6), pour 4 mL of the 60 % gradient layer solution into a centrifuge tube. Carefully overlay 4 mL of the 30 % gradient layer solution. Rest two 30/60 % gradients on ice, avoiding jolts. 3. Overlay the remaining 30/60 % gradients with 4 mL of the 18 % gradient layer solution, using a Peleus ball (see Note 6). Rest the 18/30/60 % gradients on ice, avoiding jolts.

3.2 Tissue Harvesting

1. Rats are sacrificed according to ethical guidelines and national legislation [11]. 2. Immediately rinse the organs with 10 mL ice cold 0.9 % NaCl, injecting the solution with a syringe into the portal vein (vena portae). 3. Remove liver, kidneys, heart, and brain placing them into ice cold isolation buffer with 0.1 % BSA. The heart should be placed into a small Petri dish filled with ice cold isolation buffer with 0.1 % BSA. 4. Liberate each organ from remaining fat or surrounding tissue, leaving the organs intact and rinse free from blood with ice cold isolation buffer with 0.1 % BSA. 5. Incise the heart to open the cardiac chambers and remove clotted blood if necessary. 6. Weigh liver, kidneys, heart, and brain separately (see Note 7).

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3.3 Homogenization Procedure (See Note 1) 3.3.1 Liver, Kidney, and Brain

1. Mince liver, kidneys, and brain with scissors. 2. Transfer minced liver into a 30 mL Teflon-glass homogenizer and minced kidney and brain separately into a 15 mL Teflonglass homogenizer and fill up the homogenizer with ice cold isolation buffer with 0.1 % BSA to the maximum. 3. Homogenize the tissue with 5–10 strokes at 800 rpm using the Homogenisator Potter S (see Note 8). 4. Transfer homogenized tissue to centrifuge tubes and dilute with ice cold isolation buffer with 0.1 % BSA using a total volume of ~160 mL for liver (4× 50 mL centrifugation tubes), ~40 mL for kidney (1× 50 mL centrifugation tube), and ~28 mL for brain (2× 15 mL centrifugation tubes). 5. Seal the centrifugation tubes with Parafilm® and invert.

3.3.2 Heart (See Note 9)

1. Cut the heart in ~500 mg pieces (2–3 pieces) and wash out residual blood if necessary. 2. Remove the cardiac fibrous skeleton carefully. 3. Transfer 1× 500 mg piece into a 60 mm Petri dish with 2 mL of ice cold isolation buffer with 0.1 % BSA. 4. Mince 500 mg heart tissue with scissors to obtain ~1 mm tissue pieces. 5. Mince further using a blade to obtain 0.3–0.5 mm pieces. 6. Transfer 0.3–0.5 mm heart tissue pieces into a 5 mL rough glass-in-glass homogenizer and fill up with ice cold isolation buffer with 0.1 % BSA to 5 mL. 7. Homogenize heart tissue with five strokes by hand, carefully rotating the pestle during each stroke. Transfer homogenized heart tissue to a centrifuge tube. 8. Repeat steps 3–7 until heart tissue is homogenized. 9. Dilute with ice cold isolation buffer with 0.1 % BSA using a total volume of ~30 mL (2× 15 mL centrifugation tubes). 10. Seal the centrifugation tubes with Parafilm® and invert.

3.4 Centrifugation Steps (See Note 10)

1. Centrifuge liver, kidney, brain, and heart at 800 × g for 10 min at 4 °C to free homogenates from cell debris and nuclei. 2. Transfer the supernatants to fresh centrifugation tubes. 3. Repeat steps 1 and 2 once. 4. Centrifuge liver, kidney, and heart homogenates at 9,000 × g for 10 min at 4 °C and discard the supernatants. 5. Centrifuge brain homogenate at 20,000 × g for 10 min at 4 °C and discard the supernatant (see Note 11). 6. The pellets obtained by steps 4 and 5 comprise a crude mitochondrial fraction. Resuspend this crude liver mitochondria pellet in 4–6 mL and the crude kidney mitochondria pellet in


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1 mL. Overlay 1 mL of each of the resuspended pellets on the 18 % phase of a 18/30/60 % Percoll™ gradient, using 4–6 gradients for the liver (see Note 12), and 1 gradient for the kidney. To apply Percoll™ gradient centrifugation to crude heart and brain mitochondria, resuspend each of the crude mitochondria pellets in 4 mL of 18 % gradient layer solution and overlay on the 30 % phase of a 30/60 % Percoll™ gradient. 7. Centrifuge the liver and kidney Percoll™ gradients at 9,000 × g for 10 min at 4 °C and the brain and heart Percoll™ gradients at 29,000 × g for 10 min at 4 °C. 8. Harvest the mitochondrial fraction at the boundary of the 30/60 % Percoll™ phases using a Pasteur pipette (see Note 13 and cf. Fig. 4). Transfer the mitochondrial fractions to a fresh centrifugation tube, using two tubes with a maximum capacity of ~50 mL for liver and 1 tube each for kidney, heart, and brain with a maximum capacity of 15 mL. 9. Fill all centrifugation tubes with ice cold isolation buffer without BSA to maximum capacity and centrifuge liver, kidney, heart, and brain Percoll™ gradient derived fractions at 9,000 × g for 10 min at 4 °C. 10. Discard the supernatants, resuspend the pellets and refill centrifugation tubes to maximum capacity with ice cold isolation buffer without BSA. Seal the centrifugation tubes with Parafilm® and centrifuge liver, kidney, heart, and brain Percoll™ gradient derived fractions at 9,000 × g for 10 min at 4 °C (see Note 14).

Fig. 4 Density gradient purification of mitochondrial populations using 18/30/60 % Percoll™ steps according to Subheading 3.4. The arrow indicates mitochondria gathering at the 30/60 % Percoll™ interphase. Liver and kidney mitochondria appear as prominent bands despite turbid 18 and 30 % Percoll™ phases. Heart and brain mitochondria gather in a faint but visible band. For the latter Percoll™ gradients appear with whitely clouded more intense 18 and 30 % phases than the actual mitochondria band at the 30/60 % Percoll™ interphase

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11. Discard the supernatant and resuspend the mitochondrial pellet in ice cold isolation buffer without BSA using ~3 mL for liver, ~500 μL for kidney, and 50–300 μL for heart and brain (see Note 15). Keep the isolated mitochondria on ice for further investigation (see Note 16) or freeze at −80 to −178 °C (see Note 17).

4

Notes 1. The isolation procedure is ideally performed by two persons, particularly at the homogenization steps. Recommended apportionment: person A homogenizes liver, kidney, and brain, person B takes responsibility for the homogenization of the heart. This is because heart muscle-tissue is homogenized with one additional mincing step and thereafter manually homogenized. The heart homogenization procedure takes approximately the same amount of time as the homogenization of liver, kidney, and brain together. 2. Be aware that mitochondria isolation from pancreas or spleen may require supplementation with protease inhibitors during tissue homogenization in order to retain mitochondrial functionality. 3. EGTA dissolves only slowly, causing the pH to fall to ~3. Since the supplied amount of EGTA is relatively small, care must be taken to ensure complete solubilization by visual inspection. 4. Do not re-adjust the pH once the intended pH is exceeded. 5. The indicated volumes are sufficient for up to seven Percoll™ gradients. For the liver 4–6 Percoll™ gradients are needed (see Note 12), and 1 each for kidney, heart, and brain. Prepare additional gradients if necessary. For heart and brain Percoll™ gradients prepare only a two-phase 30/60 % Percoll™ gradient. 6. The use of a Peleus ball is critical as the flow rate of a commonuse electric pipettor causes the Percoll™-layers to mix. Alternatively, a syringe can be used. Do not aim to empty the pipette (or syringe) when releasing 4 mL, as final emptying may also destroy the Percoll™ layers. 7. Weighing of organs is recommended to estimate the required number of Percoll™ gradients for the liver (see Note 12). Further, the organ weight is needed to calculate the obtained mitochondrial protein per mg organ wet weight (cf. Table 1). 8. Tissue homogenization is critical, in the sense of liberating a maximal amount of mitochondria from the cells without destroying them immediately thereafter due to excessive homogenization. Therefore, perform a minimum of strokes,


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avoiding strong pressure from the pestle to occur onto the homogenizator bottom. Liver, kidney, and brain tissues are smooth and in our experience, 5–10 strokes lead to a homogenous result in all occasions. Liver homogenate is more brownish and dense than kidney homogenate, which is a little more ginger in color. Brain homogenate is whitely beige. Observation of more reddish homogenates point to remaining blood contamination, bearing the risk of oxidative stress and is likely to worsen the functionality of isolated mitochondria. 9. Tissue homogenization is critical (see Note 8). To sufficiently homogenize the tough heart tissue, ensure the removal of the complete cardiac skeleton, as this fibrous tissue impedes mincing and fails homogenization. The mincing steps must be carried out thoroughly as heart tissue pieces greater than 0.5 mm in size also fail homogenization. During homogenization twist the pestle back and forth, avoiding strong pressure from the pestle to occur onto the homogenizator bottom. Perform a maximum of five strokes, even if some tissue pieces remain. The homogenization works best if 500 mg tissue is not exceeded. 10. For an optimal centrifugation procedure, two centrifuges are required. 11. Be careful when removing the supernatant, as the brain pellet is labile. 12. Use one Percoll™ gradient for ~2.5 g of liver wet weight. Typically 4–6 Percoll™ gradients are used for the liver. 13. A mitochondrial band is formed at the 30/60 % Percoll™ interphase. For liver and kidney, these bands are prominent, yet heart and brain mitochondrial bands appear only faint. To harvest the mitochondrial band, dip a Pasteur pipette (3 mL volume) just above the phase boundary, carefully aspirating the interphase. Immediately stop aspiration the moment the first band vanishes. Do not continue aspiration for additional overlying bands, even if they may appear denser. 14. Be careful when removing the supernatant, as the pellet may be labile. In this case Percoll™ still resides in the mitochondria pellet, making an additional washing step necessary. Decant as much supernatant as possible without losing the pellet and refill the centrifugation tube with ice cold isolation buffer without BSA to maximum capacity and repeat the centrifugation at 9,000 × g for 10 min at 4 °C. Now the brain mitochondria pellet should be sufficiently stabile to remove the supernatant completely, without pellet detachment. 15. Resuspension with the specified volumes will lead to a concentration of ~50 μg/μL liver mitochondria, 10–20 μg/μL kidney mitochondria, 3–8 μg/μL heart mitochondria, and 1–2 μg/μL brain mitochondria. Do not vortex mitochondria.

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16. Isolated mitochondria remain functional intact for at least 60 min (cf. Fig. 3). 17. Depending on the subsequent experiments, isolated mitochondria may either be frozen at −80 °C or in liquid nitrogen according to Fleischer [12], who has reported that the latter condition preserves enzymatic activity.

Acknowledgements We would like to acknowledge E.E. Rojo for critical reading of the manuscript. This study was supported in parts by the Deutsche Forschungsgemeinschaft (DFG) grant RU742/6-1 to H.Z. References 1. Löffler G, Petrides PE (1990) Biochemie und pathobiochemie, 5th edn. Springer, Berlin 2. Mootha VK et al (2003) Integrated analysis of protein composition, tissue diversity, and gene regulation in mouse mitochondria. Cell 115(5):629–640 3. Veltri KL, Espiritu M, Singh G (1990) Distinct genomic copy number in mitochondria of different mammalian organs. J Cell Physiol 143(1):160–164 4. Vijayasarathy C et al (1998) Variations in the subunit content and catalytic activity of the cytochrome c oxidase complex from different tissues and different cardiac compartments. Biochim Biophys Acta 1371(1):71–82 5. Tahara EB, Navarete FD, Kowaltowski AJ (2009) Tissue-, substrate-, and site-specific characteristics of mitochondrial reactive oxygen species generation. Free Radic Biol Med 46(9):1283–1297 6. Andreyev A, Fiskum G (1999) Calcium induced release of mitochondrial cytochrome c by different mechanisms selective for brain versus liver. Cell Death Differ 6(9): 825–832 7. Berman SB, Watkins SC, Hastings TG (2000) Quantitative biochemical and ultrastructural comparison of mitochondrial permeability transition in isolated brain and liver mitochondria: evidence for reduced sensitivity

8.

9.

10.

11.

12.

13.

of brain mitochondria. Exp Neurol 164(2): 415–425 Hogeboom GH, Schneider WC, Pallade GE (1948) Cytochemical studies of mammalian tissues; isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particulate material. J Biol Chem 172(2):619–635 Pallotti F, Lenaz G (2007) Isolation and subfractionation of mitochondria from animal cells and tissue culture lines. Methods Cell Biol 80:3–44 Petit PX et al (1998) Disruption of the outer mitochondrial membrane as a result of large amplitude swelling: the impact of irreversible permeability transition. FEBS Lett 426(1): 111–116 Close B et al (1997) Recommendations for euthanasia of experimental animals: part 2. DGXT of the European Commission. Lab Anim 31(1):1–32 Fleischer S (1979) Long-term storage of mitochondria to preserve energy-linked functions. Methods Enzymol 55:28–32 Zamzami N, Metivier D, Kroemer G (2000) Quantitation of mitochondrial transmembrane potential in cells and in isolated mitochondria. Methods Enzymol 322:208–213

14. Schmitt S et al (2014) Mitochondrion 19(Pt A): 113-123. doi:10.1016/j.mito.2014.06.005.

Chapter 8 Isolation of Mitochondria from Cultured Cells and Liver Tissue Biopsies for Molecular and Biochemical Analyses Sabine Schmitt, Carola Eberhagen, Susanne Weber, Michaela Aichler, and Hans Zischka Abstract We recently reported a new method to isolate functionally intact mitochondria from cell culture and small tissue samples (Schmitt et al., Anal Biochem 443(1):66–74, 2013). This method comprises a semiautomated cell rupture, termed pump controlled cell rupture system (PCC), which can be precisely adjusted to the specific cellular source of isolation and which can be tightly controlled (Schmitt et al., Anal Biochem 443(1):66–74, 2013). Here we provide a detailed hands-on protocol of this PCC method which results in an efficient cell breakage but preserving the mitochondrial integrity. Upon subsequent purification steps, the obtained mitochondrial fraction meets the quality and purity required for molecular analyses, e.g. proteomic comparisons, as well as for biochemical analyses, e.g. determination of diverse enzymatic activities. Key words Mitochondria, Cell culture, Biopsies, Balch homogenizer

1

Introduction Mitochondria are the cellular powerhouses and key integrators of cell death decisions [1]. Consequently, conditions that impair mitochondria can lead to severe human diseases. A prime example may be the intoxication of hepatocyte mitochondria by genetically caused excessive copper burdens in Wilson disease [2, 3]. While augmented mitochondria-dependent cell death is a major obstacle in neurodegenerative disorders [4, 5], avoidance of cell death is a hallmark of cancer [6]. Consequently, the identification of specific mitochondrial targets to either stabilize or impair their structure or function is a central aim in biomedical research [7]. Typically, the identification of such targets is achieved by comparing mitochondria isolated from healthy controls to mitochondria from diseased tissues, by proteomics, immuno-blotting or enzymatic measurements, etc. Such analyses may identify molecular and functional mitochondrial alterations that can assist in understanding disease progression [8], or in determining new targets for specific therapeutic intervention [9], or


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even provide important insights why conventional therapeutic interventions may help in some patients but fail in others [10]. Such comparative mitochondrial investigations of normal versus pathological situations require suspensions of isolated mitochondria with high and comparable purity. A most critical step in the procedure to isolate mitochondria is the initial step of cell rupture. Especially if the starting material is limited, e.g. in cases of cultured cells or tissue biopsies, an efficient cell membrane breakage is needed to obtain enough mitochondria for subsequent purification steps. However, if this is forcibly approached, a damage of the mitochondrial membranes could occur. Thus, cell rupture or tissue homogenization conditions have to be balanced between efficient plasma membrane rupture and consequent mitochondrial yield and mitochondrial integrity. To this end, means are needed that can be precisely adjusted to the respective source of isolation. We recently reported [11] that this can be accomplished using the “pump controlled cell rupture system” (PCC). PCC consists of a current version of the “Balch-homogenizer” (Isobiotec, Germany) [12] coupled to a high precision pump. Within the “Balchhomogenizer,” the cells or the tissue have to pass an exactly adjustable clearance (Fig. 1). This allows for defining and controlling the shear forces, which affect the cells and the mitochondria in the course of the homogenization process. The high precision pump further increases the reproducibility and controllability of the overall homogenization process. In order to further extend the practical value of this method, with a special focus on potential experimental pitfalls, we provide here an easy-to-follow protocol for the use of PCC and downstream centrifugation steps to isolate mitochondria from cultured cells and minute tissue samples (Fig. 2). Taken together, the herein described protocol reproducibly yields isolated mitochondria from cultured cells or minute tissue samples that meet the quality and purity required for molecular as well as for functional analyses.

2

Materials Prepare all solutions using ultrapure water (Prepared by purifying deionized water to attain a resistivity of 18.2 MΩ-cm at 25 °C).

2.1 Isolation of Mitochondria from Cell Culture or Tissue Biopsies

1. Pump controlled cell rupture system (PCC): cell homogenizer (Isobiotec, Germany), tungsten carbide balls (Isobiotec, Germany), 1 ml gastight glass syringes (internal diameter 4.608 mm, SGE Supelco, USA) (see Note 1), pump elite (Harvard apparatus, USA) (see Note 2), manual lift table. Store the manual metal-lift table at 4 °C (see Note 3). 2. Isolation buffer: 300 mM sucrose, 5 mM TES, 200 μM EGTA, pH 7.2. Transfer 102.8 g sucrose, 1.146 g TES and 76 mg

Semi-Automated Method to Isolate Mitochondria

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Fig. 1 Cartoon of the pump controlled cell rupture system (PCC). A high precision pump (1) ensures, via gastight syringes (2), the continuous sample delivery (3) in a constant rate to the “Balch-homogenizer” (4). Cell breakage occurs upon passage through a defined clearance (square), which is adjusted by selecting tungsten carbide balls of different diameters (5). The cell homogenate is collected in a second syringe (6) and can be re-subjected to the homogenizer, which is thermally equilibrated by a cooling plate (7). In this protocol, the syringe that initially contains the sample (cell suspension or tissue pieces) is referred to as syringe B and the second syringe is referred to as syringe A. Reproduced with permission from: Schmitt S, Saathoff F, Meissner L, Schropp EM, Lichtmannegger J, Schulz S, Eberhagen C, Borchard S, Aichler M, Adamski J, Plesnila N, Rothenfusser S, Kroemer G, Zischka H (2013) A semi-automated method for isolating functionally intact mitochondria from cultured cells and tissue biopsies. Anal Biochem 443:66–74

EGTA to a 1,000 ml glass beaker. Add water to a volume of 800 ml. Stir until all chemicals are dissolved and adjust pH with 5 M KOH. Transfer the buffer to a 1 L measuring cylinder, fill it up to 1,000 ml with water and store at 4 °C. The buffer can be used for 1 week.

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Fig. 2 Flow chart for the isolation of mitochondria from cultured cells and liver tissue biopsies for molecular and biochemical analyses


2.2 Purification of the Isolated Mitochondria

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1. 40 % w/v Nycodenz: transfer 40 g Nycodenz to a volumetric flask. Add 10 mM Tris–HCl (pH 7.4) to 100 ml. Store at 4 °C for up to 3 months. 2. 33 % w/v Nycodenz: Add 3.3 ml of the 40 % Nycodenzsolution and 0.7 ml isolation buffer to a 15 ml conical tube and vortex thoroughly. Store at 4 °C for no longer than 1 week. 3. 24 % w/v Nycodenz: Add 2.4 ml of the 40 % Nycodenzsolution and 1.6 ml isolation buffer to a 15 ml conical tube and vortex thoroughly. Store at 4 °C for no longer than 1 week. 4. 18 % w/v Nycodenz: Add 1.8 ml of the 40 % Nycodenzsolution and 2.2 ml isolation buffer to a 15 ml conical tube and vortex thoroughly. Store at 4 °C for no longer than 1 week. 5. Ultracentrifuge (L70, Beckman, USA), swing out rotor (SW55Ti, Beckman, USA) centrifugation tubes (Beckman Coulter, Thinwall, Ultra-Clear™, USA), 1 ml single use syringes (Omnifix®-F solo, Braun Melsungen, Germany), hypodermic needle (Sterican® 26Gx1″, Braun Melsungen, Germany).

3

Methods

3.1 Isolation of a Crude Mitochondrial Fraction from Cultured Cells and Liver Tissue

1. Pre-cool the cell homogenizer on ice for at least 10 min. Adjust the syringe pump settings concerning the syringe diameter (4.608 mm for 1 ml SGE syringe) and flow rate (700 μl/ min). Equilibrate the syringes with isolation buffer. Insert the tungsten carbide ball (see Note 4 and Table 1) into the cell homogenizer and screw it down. Flush the cell homogenizer three times with isolation buffer (see Note 5). Keep on ice until starting the homogenization. 2. For mitochondria isolation from cell culture, harvest the cells and determine the cell concentration and viability (see Note 6). Pellet the cells, discard the supernatant and resuspend the cells in isolation buffer to reach a concentration of 5 × 106 cells per ml (see Note 7). Store on ice. 3. For mitochondria isolation from liver tissue biopsy, weigh the tissue (see Note 8) and transfer it to a 10 ml glass beaker. Add 1 ml isolation buffer and scissor the liver tissue into pieces of about 1 mm3 in size. Add isolation buffer to reach a concentration of 30–40 mg of tissue per ml isolation buffer (see Note 9). Store on ice. 4. Place the cell homogenizer on the lift table. Fasten syringe A on the cell homogenizer. Fill syringe B with 1 ml of the cell suspension or of liver tissue and fasten it on the cell homogenizer.

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Table 1 Experimental settings and mitochondrial yield (9,000 × g pellet) for the isolation of mitochondria from diverse cell types (in parts reproduced with permission from Schmitt S, Saathoff F, Meissner L, Schropp EM, Lichtmannegger J, Schulz S, Eberhagen C, Borchard S, Aichler M, Adamski J, Plesnila N, Rothenfusser S, Kroemer G, Zischka H (2013) A semi-automated method for isolating functionally intact mitochondria from cultured cells and tissue biopsies. Anal Biochem 443:66–74)

Concentration [cells/ml]

Clearance [μm]

Number of strokes

Yield of crude mitochondria [μg/5 × 106 cells]

Δψ [ratio RFU ± FCCP]

Primary rat hepatocytes

3 × 106

10

3

1,230

1.8

McA 7777

5 × 106

10

3

146

1.6

5 × 10

6

6

3

82

1.6

H4IIE

7 × 10

6

6

3

70

1.9

HepT1

5 × 106

6

3

114

1.5

Huh6

5 × 106

10

3

60

1.6

HepG2

5 × 10

6

6

6

128

1.6

HEK 293

5 × 106

6

6

70

1.7

HeLa

5 × 106

6

6

107

1.7

BeWo

5 × 10

6

10

3

90

1.7

1205Lu

5 × 10

6

10

3

110

1.6

Panc02

5 × 106

6

6

162

1.5

MEF

5 × 106

6

6

90

1.6

Fao

All isolated mitochondrial suspensions efficiently built up an inner transmembrane potential (Δψm) which remained stable for at least 1 h

5. With a constant flow rate (700 μl/min) the cell suspension is pressed three or six times (see Note 10 and Table 1) through the cell homogenizer. Transfer the homogenate to a 2.0 ml tube on ice. In order to recover the whole homogenate, fill one syringe with 1 ml isolation buffer, process it once through the cell homogenizer (flow rate 700 μl/min) and transfer it also to the 2.0 ml tube (see Note 11). 6. Pass the liver tissue once (see Note 10) through the cell homogenizer (flow rate 700 μl/min) and transfer the homogenate to a 2.0 ml tube on ice. Readjust the empty syringe A on the cell homogenizer. Fill syringe B (i.e. the syringe that initially contained the tissue sample) with 1 ml isolation buffer, pump it once through the cell homogenizer (flow rate 700 μl/min) and transfer it also to the 2.0 ml tube (see Note 12). Fill syringe A with 1 ml isolation buffer and rinse the system manually (see Note 13).


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7. Repeat steps 4 and 5 or 4 and 6 until all of the cells or tissue are/is homogenized (see Note 14). 8. Centrifuge the homogenate for 5 or 10 min (cells or tissue, respectively) at 800 × g at 4 °C. 9. Transfer the supernatant to a fresh 2.0 ml tube and centrifuge it for 10 min at 9,000 × g at 4 °C. 10. Carefully discard the supernatant, slowly resuspend the pellets (crude mitochondrial fraction) in 30 μl isolation buffer. If working with multiples of 5 × 106 cells, you may pool the obtained crude mitochondrial fractions now. 11. Determine the protein concentration of the crude mitochondrial fraction. This is necessary to determine the number of Nycodenz® density gradients needed for further purification. 12. After the isolation, rinse the syringes 3–4 times with ddH2O. Clean the used tungsten carbide ball and the cell homogenizer with isopropanol and rinse them subsequently with ddH2O. Dry the tungsten carbide ball thoroughly. 3.2 Purification of the Crude Mitochondrial Fraction by Nycodenz® Density Gradient Centrifugation

1. Prepare the Nycodenz® density gradient. Add 500 μl 18 % Nycodenz® to a 5 ml centrifuge tube (Thinwall, Ultra-Clear™, Beckman Coulter, USA). Slowly syringe 300 μl of 24 or 33 % Nycodenz® (for cultured cells or liver tissue, respectively) below the 18 % layer (see Note 15). 2. Load 650–950 μg of the crude mitochondrial fraction on the discontinuous Nycodenz® density gradient (see Note 16) and centrifuge it for 15 min in an ultracentrifuge at ca. 110,000 × g (swing out rotor SW55Ti, Beckman Coulter, USA) at 4 °C. 3. Collect the mitochondria from the phase boundary (24 %/18 % or 33 %/24 % for cell culture mitochondria or rat liver tissue mitochondria, respectively) (see Note 17). Transfer the purified mitochondria into a fresh 2.0 ml tube and adjust to 2 ml with isolation buffer. Centrifuge for 10 min at 9,000 × g (4 °C) to remove Nycodenz® remnants. 4. Discard the supernatant and carefully resuspend the pellet in 30 μl isolation buffer. Purified mitochondria typically yield 20 % in mg protein of the starting amount of the crude mitochondrial isolates subjected to the Nycodenz® density gradient. 5. The purified mitochondrial fraction may now be subjected to molecular or biochemical analyses (Fig. 3 and table 1). For example upon addition of respiratory substrate (e.g. succinate) intact mitochondrial preparations should build up a membrane potential that remains stable for at least 1 h [11, 13].

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Fig. 3 Monitoring of the isolation and purification of mitochondria from H4IIE cells via a Nycodenz® density gradient (a) by immunoblot analysis (b) and electron microscopy (c)

4

Notes 1. We do not recommend using plastic syringes due to their lack of stability. In our hands, best results were obtained with syringes from the given supplier. 2. This syringe pump model can be precisely adjusted and withstands the back-pressure that emerges during the homogenization process. 3. The manual metal-lift table can thereby assist the cooling of the cell homogenizer during the homogenization process. Two manual lift tables should be at hand if more than 25 × 106 cells will be homogenized. Exchanging against a pre-cooled lift table every 30 min assists in effective cooling throughout the homogenization process.


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4. We determined a clearance of 6 or 10 μm to be successful for mitochondrial isolations from the tested cell lines given in Table 1 in terms of yield and mitochondrial integrity. We therefore recommend such initial settings which may, however, be further optimized. A clearance of 4 μm is not recommended as it impaired mitochondrial functionality. For liver tissue, a clearance smaller than 18 μm resulted in a congestion of the cell homogenizer. 5. Ensure that the system is air bubble free, as they alter the shear forces during the homogenization. Even though clearances of 6 or 8 μm already cause a noticeable back-pressure, one should be able to manually pump buffer through the cell homogenizer. If a strong resistance of the buffer flow via the PCC chamber can be encountered in such pre-isolation tests, remove the tungsten carbide ball, rinse the cell homogenizer thoroughly with isopropanol and ultrapure water and assemble the system again as described in step 1 of Subheading 3.1. 6. We typically use cell suspensions with more than 80 % viability. A lower viability might be accompanied by an impairment of mitochondrial structure or function, complicating subsequent isolation. 7. 5 × 106 cells yield crude mitochondria of at least 60 μg in the cell lines tested so far (Table 1). Less than 3 × 106 or more than 10 × 1010 cells per ml decreased either the absolute or the relative yield of isolated mitochondria, respectively. 8. The weight of tissue is required to calculate how much isolation buffer has to be added to reach a concentration of 30–40 mg tissue per ml. 9. A higher concentration overloads the system. Lower concentrations decrease the relative yield and increase the time expenditure of the homogenization. As the functional and structural stability of isolated mitochondria is time-dependent, it is advisable to minimize the endurance of the whole isolation process as much as possible. 10. The number of strokes needed to homogenize the cells depends on the respective cell line (Table 1). One “stroke” represents a passage from syringe B (i.e. the syringe that initially contains the sample) to syringe A, and vice versa. For most cell lines, less than three strokes were insufficient to break the cell membrane, whereas more than six strokes impaired mitochondrial functionality or structural integrity. For rat liver tissue, we found that more than one stroke did not markedly increase the mitochondrial yield. 11. The volume within the cell homogenizer approximately accounts for 500 μl. Therefore, it is important to rinse the system once with isolation buffer to recover the whole homogenate and to

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avoid residual contaminants in the cell homogenizer. If subsequently, mitochondria from other sources shall be isolated, we recommend at least 3–4 cleaning passages of fresh isolation buffer each. Alternatively, the whole homogenization unit may be disassembled, cleaned, and reassembled again before continuing. 12. Some small pieces of the liver tissue may enter the cell homogenizer, but not pass the clearance. We therefore recommend that the second stroke starts with the syringe used for the first stroke (syringe B). Tissue pieces that remained within the cell homogenizer can now pass the clearance with this second stroke, thereby increasing the yield without re-stressing already isolated mitochondria. 13. In order to control the number of passages for the single pieces of tissue, the system unit needs to be cleared from remaining tissue pieces before starting with the next 30–40 mg of tissue to be processed. 14. Subsequent separation of mitochondria from unbroken cells/ nuclei in a first centrifugation step at 800 × g is less efficient if the homogenate is kept on ice for more than 45 min. A plausible explanation may be a progressive clotting of mitochondria, nuclei, and cell debris over time resulting in marked mitochondrial loss in the 800 × g pellet. Thus, in the case of multiple isolations subsequent centrifugation steps should not start later than 30–40 min after the initial homogenization. 15. In order to avoid phase intermixing of the Nycodenz® density step gradient we recommend slowly layering the 300 μl of 24 or 33 % Nycodenz® (for cultured cells or liver tissue, respectively) with a syringe below the 18 % Nycodenz®. The phase boundary is slightly visible for the 18 %/24 % and clearly visible for the 18 %/33 % gradient. 16. If less than 650 μg of the crude mitochondrial fraction are loaded on the Nycodenz® gradient, the mitochondrial interphase might be barely visible, overloading (more than 950 μg of the crude mitochondrial fraction) may impair the purification efficiency. 17. To collect the mitochondria, slowly lower a pipette mounted with a 200 μl tip just above the phase boundary, carefully aspirating the interphase. Immediately stop aspiration, the moment as the mitochondrial band vanishes.

Acknowledgement We would like to acknowledge E.E. Rojo and A. Simmons for critical reading of the manuscript. This study was supported in parts by the Deutsche Forschungsgemeinschaft (DFG) grant RU742/6-1 to H.Z.


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References 1. Green DR, Kroemer G (2004) The pathophysiology of mitochondrial cell death. Science 305(5684):626–629 2. Zischka H, Lichtmannegger J (2014) Pathological mitochondrial copper overload in livers of Wilson’s disease patients and related animal models. Ann N Y Acad Sci 1315:6–15 3. Zischka H et al (2011) Liver mitochondrial membrane crosslinking and destruction in a rat model of Wilson disease. J Clin Invest 121(4):1508–1518 4. Lin MT, Beal MF (2006) Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature 443(7113):787–795 5. Winklhofer KF, Haass C (2010) Mitochondrial dysfunction in Parkinson’s disease. Biochim Biophys Acta 1802(1):29–44 6. Hanahan D, Weinberg RA (2000) The hallmarks of cancer. Cell 100(1):57–70 7. Schmitt S et al (2014) Why to compare absolute numbers of mitochondria. Mitochondrion 19(A):113–123 8. Schulz S et al (2013) Progressive stages of mitochondrial destruction caused by cell toxic

9.

10.

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bile salts. Biochim Biophys Acta 1828(9): 2121–2133 Galluzzi L et al (2010) Mitochondrial gateways to cancer. Mol Aspects Med 31(1): 1–20 Aichler M et al (2013) Clinical response to chemotherapy in oesophageal adenocarcinoma patients is linked to defects in mitochondria. J Pathol 230(4):410–419 Schmitt S et al (2013) A semi-automated method for isolating functionally intact mitochondria from cultured cells and tissue biopsies. Anal Biochem 443(1):66–74 Balch WE, Rothman JE (1985) Characterization of protein transport between successive compartments of the Golgi apparatus: asymmetric properties of donor and acceptor activities in a cell-free system. Arch Biochem Biophys 240(1):413–425 Zamzami N, Metivier D, Kroemer G (2000) Quantitation of mitochondrial transmembrane potential in cells and in isolated mitochondria. Methods Enzymol 322:208–213

Chapter 9 Dynamic Range Compression with ProteoMiner™: Principles and Examples Lei Li Abstract One of the main challenges in proteomics investigation, protein biomarker research, and protein purity and contamination analysis is how to efficiently enrich and detect low-abundance proteins in biological samples. One approach that makes the detection of rare species possible is the treatment of biological samples with solid-phase combinatorial peptide ligand libraries, ProteoMiner. This method utilizes hexapeptide bead library with huge diversity to bind and enrich low-abundance proteins but remove most of the high-abundance proteins, therefore compresses the protein abundance range in the samples. This work describes optimized protocols and highlights on the successful application of ProteoMiner to protein identification and analysis. Key words Low-abundance proteins, Proteomics, ProteoMiner, Dynamic range compression, Plasma, Serum, Saliva, Host cell proteins

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Introduction Concentrations of different proteins in biological samples can span as much as 12 orders of magnitudes. For example, 99 % of the protein content in plasma and serum samples is only about 20 highabundance proteins, whereas the other thousands of proteins are less than 1 % of the plasma or serum proteome [1, 2]. After decades of research and accumulated knowledge on high-abundance proteins due to their relatively easy access, the discovery and analysis of the rare species in proteomes attracts more and more attentions. ProteoMiner technology is based on a combinatorial hexapeptide library bound to chromatographic beads [3–6]. It can be used to decrease levels of high-abundance proteins for researchers to enrich and detect more mid- to low-abundance proteins in complex samples and unveil the proteomes deeply and comprehensively. When a complex biological sample is added to the beads, proteins bind to their specific ligands through combination of various types of interactions including ionic interaction, hydrophobic interaction, hydrogen bonding and van


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Fig. 1 (a) Scheme of the ProteoMiner technology to remove most of high-abundant proteins and enrich lowabundant proteins, thus compressing the protein concentration range in biological samples. (b) Twodimensional gel electrophoresis of human serum samples before and after treatment by ProteoMiner beads, adapted from [31]. The samples were first separated on a pH 5–8 IPG strip, then an 8–16 % Criterion Tris–HCl gel before visualization by the Flamingo Fluorescent Gel Stain

der Waal’s force, so high-abundance proteins saturate their specific ligands and the excess proteins are washed away. In contrast, mediumand low-abundance proteins not saturating their specific ligands will mostly stay bound to their peptide ligands and are therefore enriched relatively to the high-abundance proteins during the process (Fig. 1). The result is a reduction in the dynamic range which allows for the detection of medium- and low-abundance proteins. The ProteoMiner technology has been extensively applied to variety of sample types [6] including plasma and serum [3], urine [7], bile [8], platelets [9], red blood cell extract [10, 11], egg white extract [12], egg yolk extract [13], CSF [14], saliva [15, 16], venom [17], milk whey fraction [18, 19], sea urchin coelomic fluid [20], plant leaf [21], seeds [22], peel and pulp [23], phloem exudates [24, 25], plant latex [26], bacteria [3], and even beverage and wine [27, 28]. The ProteoMiner beads have also been successfully used on detection, identification, adsorption, and removal of impurities or host cell proteins from purified proteins [29, 30]. Even though ProteoMiner beads have a large diversity of hexapeptides, the analysis in different groups [31, 32] using twodimensional gel electrophoresis, immunoassay and mass spectrometry, indicated a high level of consistency from sample to sample when processed with similar and variable bead volumes.

Principles and Examples of ProteoMiner

200 µl

500 µl

1 ml

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5 ml 250 kD 150 kD 100 kD 75 kD 50 kD 37 kD 25 kD 20 kD 15 kD

Fig. 2 Effect of sample to beads ratio on protein enrichment profile. Different amount of human serum samples were treated with 20 μl ProteoMiner beads in triplicate. The eluates were separated on a 4–20 % criterion Tris–HCl gel and visualized by Bio-Safe Coomassie Stain. The red arrows indicate increasing amount of proteins with higher sample to beads ratio

This chapter describes the standard protocol optimized for plasma and serum samples to compress proteomic dynamic range. Results with other sample types will vary depending on the amount of protein in the samples as well as the dynamic range that each sample type contains. The ratio of protein to beads is crucial for optimal performance of ProteoMiner kits, so best results will be obtained by optimization of ratio of proteins to ProteoMiner beads for each individual sample type. With increasing amount of serum sample loaded onto same amount of ProteoMiner beads, some proteins can be enriched more than the others (Fig. 2).

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Materials 1. Microcentrifuge or vacuum manifold. 2. ProteoMiner Protein Enrichment Large Capacity Kit (BioRad, #163-3007 or #163-3009) and Small Capacity Kit (BioRad, #163-3006 or #163-3008). These kits contain 100 μl (Large Capacity) or 20 μl (Small Capacity) of ProteoMiner beads, PBS wash buffer (150 mM NaCl, 10 mM NaH2PO4, pH 7.4), lyophilized elution reagent (8 M urea, 2 % CHAPS) and rehydration reagent (5 % acetic acid), and are intended for simple one-step elution of the bound proteins. 3. ProteoMiner Sequential Elution Large Capacity Kit (Bio-Rad, #163-3011) and Small Capacity Kit (Bio-Rad, #163-3010). These kits contain 100 μl (Large Capacity) or 20 μl (Small Capacity) of ProteoMiner beads, PBS wash buffer (150 mM NaCl, 10 mM NaH2PO4, pH 7.4) and 4 elution reagents

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(Elution Reagent 1: 1 M NaCl, 20 mM HEPES, pH 7.4; Elution Reagent 2: 200 mM glycine, pH 2.4; Elution Reagent 3: 60 % ethylene glycol in water; Elution Reagent 4: 33.3 % 2-propanol, 16.7 % acetonitrile, 0.1 % trifluoroacetic acid), and are intended for sequential elution of the bound proteins. 4. Micro Bio-Spin P-6 Gel Columns in 10 mM Tris–HCl buffer, pH 7.4 (Bio-Rad, #732-6221 or #732-6222). 5. Exchange buffer: 7 M Urea, 2 M thiourea, 2 % CHAPS with or without 30 mM Tris–HCl, pH 8.5. 6. 2 ml microcentrifuge tubes with and without caps.

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Methods For Subheadings 3.1–3.5, the ProteoMiner Large Capacity Kits are used as the example. The bold and italic volumes in parenthesis are for the ProteoMiner Small Capacity Kits.

3.1 Column Preparation and Equilibration

Vacuum at about 16 mmHg can replace centrifugation for column preparation, sample binding, and sample wash steps if desired. 1. First remove the top cap and then snap off the bottom cap from each of the ProteoMiner spin columns you will be using. Keep both top and bottom caps for usage throughout the protocol. If beads settle in top cap, replace after removing bottom plug and centrifuge with top cap on column. To use bottom cap as a plug, invert and firmly place to bottom of spin column. 2. Place the column in a collection tube and centrifuge at 1,000 × g for 30–60 s to remove the storage solution. 3. Replace the bottom cap and add 600 μl (200 μl) of wash buffer, then replace top cap. Invert column end-to-end several times over a 5 min period. 4. Remove bottom cap, place the column in a collection tube and centrifuge at 1,000 × g for 30–60 s to remove the wash buffer. 5. Repeat steps 3 and 4 one more time.

3.2 Sample Binding (See Note 1)

Samples should be free of precipitate. If needed, centrifuge samples at 10,000 × g for 10 min to clarify or pretreat the samples to remove incompatible or interfering materials (see Note 2). Replace bottom cap and add sample to column, for example 1 ml (200 μl) of serum or plasma. Replace top cap and invert column end-to-end on a platform or rotational shaker for 2 h at room temperature (see Note 3).


3.3

Sample Wash

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1. Remove bottom cap, place column in a collection tube and centrifuge at 1,000 × g for 30–60 s. 2. Replace bottom cap and add 600 μl (200 μl) of wash buffer to column. Replace top cap and invert from end-to-end over a 5 min period. 3. Remove bottom cap, place column in a collection tube and centrifuge at 1,000 × g for 30–60 s. 4. Repeat steps 2 and 3 three more times.

3.4

Elution

1. After all wash buffers have been removed, replace the bottom and bottom caps and add 600 μl (200 μl) deionized water. Attach top cap and invert end-to-end for 1 min. 2. Remove caps, place column in a collection tube and centrifuge at 1,000 × g for 30–60 s to remove water. If using vacuum up to this point, you will now need to switch to centrifugation. 3. Attach bottom cap to the column (take caution to ensure the bottom cap is tightly attached). Add 100 μl (20 μl) of rehydrated elution reagent to the column and replace top cap. Lightly vortex for 5 s (see Note 4). 4. Incubate column at room temperature, lightly vortex several times over a period of 15 min. 5. Remove caps, place in a clean collection tube and centrifuge at 1,000 × g for 30–60 s. This elution contains your eluted proteins. 6. Repeat steps 5–7 two more times. 7. Store elution at −20 °C or proceed with downstream analysis. For some downstream applications, we recommend a clean-up of your sample prior to analysis. See Subheading 3.6 below for the protocol in details.

3.5 Sequential Elution (Alternative to Subheading 3.4 After Subheading 3.3)

1. Carefully add 200 μl wash buffer on top cap and all sides of the column to ensure none of the beads are stuck to the cap and sides of the column. 2. Centrifuge at 1,000 × g for 30–60 s. Discard collected material. If using vacuum up to this point, you will now need to switch to centrifugation. 3. Attach bottom cap to the column (take caution to ensure the bottom cap is tightly attached). Add 100 μl (20 μl) of elution reagent 1 to spin column. Incubate at room temperature and lightly vortex several times over a period of 10 min. 4. Remove bottom cap, place column in a collection tube and centrifuge at 1,000 × g for 30–60 s to collect the elution. This elution contains your eluted proteins. 5. Repeat steps 3 and 4 two more times and collect both elutions in the same tube from the above step.

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6. Repeat steps 3–5 for the other three elution reagents. 7. Store elutions at −20 °C or proceed with downstream analysis. 3.6 Elution Clean-Up with Micro Bio-Spin columns

Some downstream applications of the ProteoMiner-treated elutions may require to remove incompatible reagents. Below is an easy and convenient method using the Micro Bio-Spin P-6 gel column for buffer exchanging the ProteoMiner eluates with compatible reagents. 1. Invert the column several times to resuspend the settled gel and remove any bubbles. 2. Snap off the tip and place column in a 2 ml microcentrifuge tube. Remove cap. Allow the excess packing buffer to drain by gravity to top of gel bed. (If column does not begin to flow, push cap back into column and remove.) Discard the buffer. 3. Place the column back into the 2 ml tube. Centrifuge for 1 min at 1,000 × g to remove the packing buffer. 4. Block the bottom of the Bio-Spin 6 column with the tip and add 500 μl of the downstream application compatible buffer (for example, the Exchange buffer in Subheading 2). Invert the column several times to resuspend the settled gel. 5. Remove the bottom tip and place the column in a clean 2 ml microcentrifuge tube. Remove cap. Allow the excess packing buffer to drain by gravity to top of gel bed. Discard the buffer. 6. Place the column back into the 2 ml tube. Centrifuge for 1 min at 1,000 × g to remove the buffer. 7. Repeat steps 4–6 two more times. 8. Place the column in a clean 2 ml microcentrifuge tube. Carefully apply up to 50 μl of ProteoMiner eluates to the column and centrifuge for 2 min at 1,000 × g. The collected solution is buffer-exchanged to the compatible reagents.

4

Notes 1. As listed examples in the introduction, variety of sample types from different species and sources have been used successfully with the ProteoMiner beads. In principle, the more proteins are incubated with the ProteoMiner beads, the more degree of dynamic range compression can be reached, whereas extremely low abundant proteins can be much enriched compared to the high abundant proteins. As a guideline, 50 mg of serum or plasma proteins are recommended for 100 μl ProteoMiner beads. In return, a bit more than 1 mg of proteins can be bound to and eluted from the beads, so that up to 50-fold dynamic range compression can be obtained. Depending on the needs and goals of the specific research, it would be wise to keep in


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mind of this while designing the experiment and deciding the ratio between sample and ProteoMiner beads. Much higher sample to beads ratio has been tested (Fig. 2) with still increasing enrichment of a few proteins. Regular buffers based on phosphate buffered saline (PBS) or Tris buffered saline (TBS) with or without mild detergents in samples are recommended for the ProteoMiner beads because of their less interference with binding of proteins to ProteoMiner peptides (see Notes 2–4 below in more details). 2. Biological samples often contain interfering or incompatible materials other than proteins which researchers are interested in. It is crucial to remove them before applying samples onto ProteoMiner beads. Dialysis, gel filtration, centrifugation and protein precipitation are a few examples of pretreatment methods, and can be selected according to the sample types and sizes, protein concentrations, and downstream applications. The quality of the samples before ProteoMiner treatment can be simply analyzed by gel electrophoresis or a UV/Vis spectrophotometer. The ProteoMiner process can be viewed as affinity purification with complex protein–peptide interactions. Interfering materials and incompatible solutions for regular protein affinity purification will have some effects on binding of certain proteins to their specific peptide ligands. Buffer compatibility for wash and elution steps can often refer affinity purification guidance and principles. 3. It is recommended to optimize the sample to beads ratio in order to achieve the desired performance. Higher sample to beads ratio tends to enrich more mid- to low-abundant proteins (Fig. 2). If the protein concentration is low in the samples, concentrating may be needed. Otherwise, the equilibrated ProteoMiner beads can be taken out of the column and incubated with samples in another tube. After sample binding is done, the beads can be spun down and transferred back into the column for wash and elution. The sample binding incubation time and temperature in this step are the recommendations for plasma and serum samples. Incubation at 4 °C overnight would be another way researchers can often use for their specific samples. If using plasma, clumping may occur after 1 h of binding; this is expected and will not negatively impact the sample preparation. Heparinized plasma is not compatible with this kit. 4. For 2-D users who plan to use the ProteoMiner treated samples on DIGE, this elution reagent will require clean up and pH adjustment. As an alternative you may elute with DIGE buffer. However, this may result in a decreased yield and number of protein spots. For a more complete elution if it would not interfere with downstream applications, the Laemmli sample buffer with 1 % SDS and reducing agents like β-Mercaptoethanol or DTT can be added and incubated with ProteoMiner beads at 95 °C for 5 min before elution.

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References 1. Anderson N, Anderson N (2002) The human plasma proteome: history, character, and diagnostic prospects. Mol Cell Proteomics 11: 845–867 2. Anderson N, Polanski M, Pieper R et al (2004) The human plasma proteome: a nonredundant list developed by combination of four separate sources. Mol Cell Proteomics 3:311–326 3. Thulasiraman V, Lin S, Gheorghiu L et al (2005) Reduction of the concentration difference of proteins in biological liquids using a library of combinatorial ligands. Electrophoresis 26:3561–3571 4. Guerrier L, Thulasiraman V, Castagna A et al (2006) Reducing protein concentration range of biological samples using solid-phase ligand libraries. J Chromatogr B 833:33–40 5. Boschetti E, Righetti P (2008) Hexapeptide combinatorial ligand libraries: the march for the detection of the low-abundance proteome continues. Biotechniques 44:663–665 6. Boschetti E, Righetti P (2013) Low abundance protein discovery: state of the art and protocols. Elsevier, Amsterdam 7. Castagna A, Cecconi D, Sennels L et al (2005) Exploring the hidden human urinary proteome via ligand library beads. J Proteome Res 4:1917–1930 8. Guerrier L, Claverol S, Finzi L et al (2007) Contribution of solid-phase hexapeptide ligand libraries to the repertoire of human bile proteins. J Chromatogr A 1176:192–205 9. Guerrier L, Claverol S, Fortis F et al (2007) Exploring the platelet proteome via combinatorial, hexapeptide ligand libraries. J Proteome Res 6:4290–4303 10. Simó C, Bachi A, Cattaneo A et al (2008) Performance of combinatorial peptide libraries in capturing the low-abundance proteome of red blood cells. 1. Behavior of mono- to hexapeptides. Anal Chem 80:3547–3556 11. Bachi A, Simó C, Restuccia U et al (2008) Performance of combinatorial peptide libraries in capturing the low-abundance proteome of red blood cells. 2. Behavior of resins containing individual amino acids. Anal Chem 80: 3557–3565 12. D'Ambrosio C, Arena S, Scaloni A et al (2008) Exploring the chicken egg white proteome with combinatorial peptide ligand libraries. J Proteome Res 7:3461–3474 13. Farinazzo A, Restuccia U, Bachi A et al (2009) Chicken egg yolk cytoplasmic proteome, mined via combinatorial peptide ligand libraries. J Chromatogr A 1216:1241–1252

14. Shores K, Udugamasooriva D, Kodadek T et al (2008) Use of peptide analogue diversity library beads for increased depth of proteomic analysis: application to cerebrospinal fluid. J Proteome Res 7:1922–1931 15. Bandhakavi S, Stone M, Onsongo G et al (2009) A dynamic range compression and three-dimensional peptide fractionation analysis platform expands proteome coverage and the diagnostic potential of whole saliva. J Proteome Res 8:5590–5600 16. Bandhakavi S, Van Riper S, Tawfik P et al (2010) Hexapeptide libraries for enhanced protein PTM identification and relative abundance profiling in whole human saliva. J Proteome Res 10:1052–1061 17. Calvete J, Fasoli E, Sanz L et al (2009) Exploring the venom proteome of the western diamondback rattlesnake, Crotalus atrox, via snake venomics and combinatorial peptide ligand library approaches. J Proteome Res 8:3055–3067 18. D'Amato A, Bachi A, Fasoli E et al (2009) In-depth exploration of cow’s whey proteome via combinatorial peptide ligand libraries. J Proteome Res 8:3925–3936 19. Liao Y, Alvarado R, Phinney B et al (2011) Proteomic characterization of human milk whey proteins during a twelve-month lactation period. J Proteome Res 10:1746–1754 20. Fasoli E, D'Amato A, Righetti P et al (2012) Exploration of the sea urchin coelomic fluid via combinatorial peptide ligand libraries. Biol Bull 222:93–104 21. Fasoli E, D’Amato A, Kravchuk A et al (2011) Popeye strikes again: the deep proteome of spinach leaves. J Proteomics 74:127–136 22. Esteve C, D'Amato A, Marina M et al (2012) Identification of olive (Olea europaea) seed and pulp proteins by nLC-MS/MS via combinatorial peptide ligand libraries. J Proteomics 75:2396–2403 23. Fasoli E, Righetti P (2013) The peel and pulp of mango fruit: a proteomic samba. Biochim Biophys Acta 1834:2539–2545 24. Fröhlich A, Gaupels F, Sarioglu H et al (2012) Looking deep inside: detection of lowabundance proteins in leaf extracts of Arabidopsis and phloem exudates of pumpkin. Plant Physiol 159:902–914 25. Gaupels F, Sarioglu H, Beckmann M et al (2012) Deciphering systemic wound responses of the pumpkin extrafascicular phloem by metabolomics and stable isotope-coded protein labeling. Plant Physiol 160:2285–2299

Principles and Examples of ProteoMiner 26. D’Amato A, Bachi A, Fasoli E et al (2010) In-depth exploration of Hevea brasiliensis latex proteome and “hidden allergens” via combinatorial peptide ligand libraries. J Proteomics 73:1368–1380 27. Cereda A, Kravchuk A, D’Amato A et al (2010) Proteomics of wine additives: mining for the invisible via combinatorial peptide ligand libraries. J Proteomics 73:1732–1739 28. Cereda A, Kravchuk A, D’Amato A et al (2010) Noah’s nectar: the proteome content of a glass of red wine. J Proteomics 73:2370–2377 29. Fortis F, Guerrier L, Righetti P et al (2006) A new approach for the removal of protein impurities from purified biologicals using

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combinatorial solid-phase ligand libraries. Electrophoresis 27:3018–3027 30. Fortis F, Guerrier L, Areces L et al (2006) A new approach for the detection and identification of protein impurities using combinatorial solid phase ligand libraries. J Proteome Res 5:2577–2585 31. Li L, Sun C, Freeby S et al (2009) Protein sample treatment with peptide ligand library: coverage and consistency. J Proteomics Bioinform 2:485–494 32. Dwivedi R, Krokhin O, Cortens J et al (2009) Assessment of the reproducibility of random hexapeptide peptide library-based protein normalization. J Proteome Res 9:1144–1149

Chapter 10 Qualitative and Quantitative Proteomic Analysis of Formalin-Fixed Paraffin-Embedded (FFPE) Tissue Omid Azimzadeh, Michael J. Atkinson, and Soile Tapio Abstract Formalin-fixed, paraffin-embedded (FFPE) tissue has recently gained interest as an alternative to fresh/ frozen tissue for retrospective protein biomarker discovery. However, during the formalin fixation proteins undergo degradation and cross-linking, making conventional protein analysis technologies challenging. Cross-linking is even more challenging when quantitative proteome analysis of FFPE tissue is planned. The use of conventional protein labeling technologies on FFPE tissue has turned out to be problematic as the lysine residue labeling targets are frequently blocked by the formalin treatment. We have established a qualitative and quantitative proteomics analysis technique for FFPE tissues that combines label-free proteomic analysis with optimized protein extraction and separation conditions. Key words Label-free proteomics, Biomarker discovery, Mass spectrometry, Formalin-fixed paraffinembedded (FFPE), Peptide modification

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Introduction For decades, formalin-fixed and paraffin-embedded (FFPE) tissue has been the standard for histopathological analysis due to the ease of sample preparation and stability in long-term storage. Clinical archives frequently combine stored FFPE samples with data on diagnosis and outcome. Such samples hold unique information on biological pathways and cellular processes leading to disease, but suitable molecular technologies are required to access the information [1–3]. For a long time, quantitative proteomic studies on archival material have been considered to be an almost impossible task, primarily due to the harsh and irreversible procedures needed for fixation and the loss of macromolecular integrity during prolonged storage. The changes occurring during cross-linking of proteins during the fixation of FFPE tissue have been investigated in different studies [4–6]. The consensus is that the major consequence of formaldehyde fixation is the generation of a methylol modification at free lysine residues [4, 7]. Several studies have been conducted to evaluate different components


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used for protein extraction, including buffer components, detergents, pH, temperature, and pressure [4, 8–10]. Most chemical labels used in quantitative proteomics target lysine residues, leading to inefficient labeling of FFPE material where lysine is no longer available [11–13]. Recently, label-free approaches have been proposed as an alternative for quantification of proteome profiles from FFPE samples [14, 15]. Here, we provide a tested methodological protocol suitable for the quantitative and qualitative proteomic analysis of archival tissue combining optimal protein extraction and separation with gelbased proteomics [4]. We successfully used this method to study the FFPE archived cardiac tissue from sham- and total bodyirradiated C57BL/6 mice [4, 16]. For quantitative proteome analysis, we used label-free approach [17] to detect putative molecular biomarkers of ionizing radiation. This methodology can be transferred to other tissues and treatments. The protocol will facilitate the development of future proteomic analysis techniques for FFPE tissue and provide a tool for the validation in archival samples of biomarkers of exposure, prognosis, and disease.

2

Materials All solutions are prepared using HPLC grade water. All buffers contain protease inhibitor cocktail following the manufacturer’s instructions (Complete, Roche Diagnostics).

2.1 Buffers and Solutions for Sample Preparation

1. Rehydration buffer: Graded series of ethanol (100, 95, and 70 %) (v/v) in water. 2. Wash buffer: 20 mM Tris–HCl, pH 7.5, 0.5 % (w/v) beta-octylglucoside. 3. Lysis buffer: 20 mM Tris–HCl, pH 8.8, 2 % (w/v) SDS, 1 % (w/v) beta-octylglucoside, 200 mM DTT, 200 mM glycine.

2.2 SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE)

1. 2× Laemmli sample buffer: 62.5 mM Tris–HCl, pH 6.8, 2 % (w/v) SDS, 25 % (w/v) glycerol, 0.01 % (w/v) bromophenol blue, 5 % β-mercaptoethanol. 2. Homogeneous pre-cast SDS-PAGE gels (12 %). 3. SDS-PAGE running buffer: 192 mM glycine, 0.1 % SDS (w/v), 25 mM Tris base. 4. Electrophoresis equipment: running chamber and power supply. 5. Colloidal Coomassie Blue staining buffer: 0.08 % (w/v) Coomassie Blue G250 (CBB G250), 1.6 % (v/v) orthophosphoric acid, 8 % (w/v) ammonium sulfate, and 20 % (v/v) methanol.

FFPE Qualitative and Quantitative Proteomics

2.3 Processing of Gel-Resolved Proteins and Tryptic Digestion

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1. Trypsin (Sigma-Aldrich). 2. 100 % acetonitrile (ACN). 3. 50 mM NH4HCO3 in 30 % ACN (w/v), pH 8.0. 4. 10 mM NH4HCO3, pH 8.0. 5. 80 % (v/v) ACN, 1 % (v/v) trifluoroacetic acid (TFA). 6. 5 % (v/v) ACN, 0.5 % (v/v) TFA. 7. Trypsin digestion equipment: shaker, incubator, and SpeedVac centrifuge. 8. Peptide separation and identification equipment: Highperformance liquid chromatography (HPLC) and electrospray ionization tandem mass spectrometry (ESI-MS/MS).

2.4 Experimental Kits

1. 2D-Clean-Up-Kit (Bio-Rad).

2.5

1. Progenesis software (Nonlinear).

Software

2. Bradford reagent.

2. Mascot (Matrix Science). 3. PANTHER bioinformatics tool (Protein ANalysis THrough Evolutionary Relationships). 4. Database for Annotation, Visualization, and Integrated Discovery (DAVID).

3

Methods

3.1 Protein Extraction from FFPE Tissue

1. Cut 20 μm sections from the FFPE tissue blocks after initial trimming to remove air exposed surfaces (see Note 1). 2. Place FFPE tissue sections (20 μm thick, 80 mm2 wide) on microscope slides and deparaffinize by incubating twice with xylene for 10 min at room temperature before rehydration in a graded series of ethanol (100, 95, and 70 %) for 10 min each. 3. Scrap the tissue sections from the slides and transfer to a reaction cup (see Note 1). 4. Wash the tissue sections with wash buffer described in Subheading 2.1 for 15 min at room temperature with slight shaking. 5. Incubate all samples in lysis buffer at 100 °C for 20 min, and then at 80 °C for 2 h with shaking (see Note 2). 6. Centrifuge the extracts for 30 min at 14,000 × g at 4 °C. 7. Precipitate the protein extract with the 2D clean-up kit following the manufacturer’s instructions (see Note 3). 8. Estimate the protein concentration in the lysate by the Bradford assay (see Notes 4 and 5).

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9. Dissolve protein pellet (see step 7) with 1× Laemmli buffer at a concentration of 1–5 μg/μL. 3.2 SDS-PAGE and Protein Staining

1. Load sample wells of the 4–12 % SDS-PAGE gel with protein solution and separate equal amounts (50–100 μg protein per lane) [18] (see Note 6). 2. Fill electrophoresis chamber with SDS-PAGE running buffer. Run electrophoresis gels at 90 V for the first ~15 min at room temperature (until the blue dye moved out of the stacking gel). Thereafter, increase voltage to 120 V. 3. After electrophoresis incubate 1D PAGE gels with Colloidal Coomassie solution overnight on a shaker at 4 °C. 4. Destain the gels with water and transfer to a clean container and store in water at 4 °C until further analysis.

3.3 Processing of Gel-Resolved Proteins and Tryptic Digestion

For the identification and quantification of proteins, slice each SDS-PAGE lane horizontally into five pieces. Digest proteins into peptides prior to mass spectrometry analysis as follows: 1. Destain the gel pieces further and rinse with buffer containing 50 mM NH4HCO3 in 30 % acetonitrile (ACN) (v/w), pH 8.0. 2. Equilibrate the gel pieces in 10 mM NH4HCO3, pH 8.0, prior to proteolytic digestion by shrinking them in 100 % ACN and rehydrating in 10 mM NH4HCO3. 3. Add 0.1–0.2 μg of trypsin per gel piece and incubate overnight at 37 °C (see Note 7). 4. Elute the digested proteins using 80 % (v/v) ACN, 1 % (v/v) trifluoroacetic acid (TFA). 5. Dry the eluates in a SpeedVac centrifuge. 6. Resuspend the dried samples in 20 μl 5 % (v/v) ACN, 0.5 % (v/v) TFA for subsequent high-performance liquid chromatography (HPLC) separation and ESI-MS/MS analysis (see Notes 8–10).

3.4 An Example of Label-Free Proteomics on FFPE Sample

Due to the protein cross-linking during FFPE preparation, conventional labeling approach cannot be used for quantitative proteomic analysis on FFPE tissue. Formaldehyde, used in tissue fixation, leads to a 30 Da (methylol) modification mainly on lysine residues and inhibits classical labeling methodology that is targeted to lysine residues. Alternatively, FFPE proteome profile has been recently quantified using non-labeling approaches. For label-free proteomics, the acquired spectra should be identified and quantified using the Progenesis software [16, 17] (see Note 11). To interpret the observed alterations in proteome, the significantly differentially expressed proteins in all samples should be analyzed using bioinformatics tools such as PANTHER or DAVID [19, 20]. Classify the proteins using Gene ontology (GO) categories [21].


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The described methodology has been used to study the FFPE archived cardiac tissue from sham- and total body-irradiated C57BL/6 mice [4, 16]. The samples were analyzed using labelfree approach to detect putative molecular biomarkers of ionizing radiation. The differentially expressed proteins were classified using GO categories (cellular components, molecular function, and biological processes). Differentially expressed proteins grouped according to molecular functions are shown in Fig. 1. In our case, proteins with oxidoreductase activity represented the largest group. The GO analysis of biological processes indicated that the deregulated proteins were mainly involved in lipid, carbohydrate, phosphate and nucleic acid metabolism as well as molecular transport (Fig. 1b). The GO cellular compartment analysis showed that most deregulated proteins belonged to the mitochondrial proteome. Functional annotation clustering analysis using the DAVID software showed that the proteins belonged to different mitochondrial compartments such as membranes, electron transport machinery, matrix, and channels (Fig. 1c). Furthermore, radiation-induced

Fig. 1 Radiation-induced alteration of the cardiac mitochondrial proteome. Significantly differentially regulated proteins were classified for GO categories “Molecular function” (a), “Biological process” (b), and “Cellular compartment” (c) using the PANTHER and DAVID bioinformatics tools. The amount of deregulated proteins is indicated as a number of the total amount of up- or down-regulated proteins at 24 h. The analysis indicated that ionizing radiation caused impairment of mitochondrial structure and function

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changes affected the proteins involved in oxidative phosphorylation such as NADH dehydrogenase, cytochrome c, and succinate dehydrogenase.

4

Notes 1. To avoid keratin contamination, always wear gloves and cover the hair during handling samples and gels. 2. To minimize proteolysis of the protein lysates, it is recommended to use protease inhibitors and keep the samples on ice. 3. During precipitation, do not overdry the pellet after removal of the acetone as this might lead to difficulty in resolubilization of the pellet in the resuspension buffer. 4. If you have problems in determining the protein concentration of the lysate, it may be due to some component in the lysis buffer. In this case try another protein quantification method. 5. If the protein yield after resolubilization is low, test other solubilization buffers as the result may differ with the types of tissues used. For more information, see the protocols mentioned in ref. 6. 6. Use ultrapure proteomics grade reagents to prepare buffers for staining and digestion. 7. For peptide or protein digestion, a ratio of between 1:100 and 1:20 (w/w) of trypsin to substrate is recommended. 8. HPLC setting and proteomics quantification should be handled by trained researchers. 9. To achieve a comprehensive and statically significant interpretation, use at least five biological and technical replicates if they are available. 10. In order to evaluate the technical variability of mass spectrometry runs of label-free peptide quantifications of FFPE samples, analyze the aliquot of one control sample by repetitive LC-MS/MS runs at least three replicates. 11. In order to evaluate the degree of the reversion of the crosslinking, the variable 30 Da methylol modification caused by the cross-linking [7, 22, 23] was set for lysine, histidine, and arginine residues during searching against the database [4].

Acknowledgements This work was supported by a grant from the European Community's Seventh Framework Programme (EURATOM) contract n°232628 (STORE).


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References 1. Tapio S, Atkinson MJ (2008) Molecular information obtained from radiobiological tissue archives: achievements of the past and visions of the future. Radiat Environ Biophys 47(2): 183–187 2. Tapio S, Hornhardt S, Gomolka M et al (2010) Use of proteomics in radiobiological research: current state of the art. Radiat Environ Biophys 49(1):1–4 3. Tapio S, Schofield PN, Adelmann C et al (2008) Progress in updating the European radiobiology archives. Int J Radiat Biol 84(11):930–936 4. Azimzadeh O, Barjaktarovic Z, Aubele M et al (2010) Formalin-fixed paraffin-embedded (FFPE) proteome analysis using gel-free and gel-based proteomics. J Proteome Res 9(9): 4710–4720 5. Giusti L, Lucacchini A (2013) Proteomic studies of formalin-fixed paraffin-embedded tissues. Expert Rev Proteomics 10(2):165–177 6. Steiner C, Ducret A, Tille JC et al (2014) Applications of mass spectrometry for quantitative protein analysis in formalin-fixed paraffin-embedded tissues. Proteomics 14(4–5): 441–451 7. Metz B, Kersten GF, Hoogerhout P et al (2004) Identification of formaldehydeinduced modifications in proteins: reactions with model peptides. J Biol Chem 279(8): 6235–6243 8. Magdeldin S, Yamamoto T (2012) Toward deciphering proteomes of formalin-fixed paraffin-embedded (FFPE) tissues. Proteomics 12(7):1045–1058 9. Hatakeyama K, Wakabayashi-Nakao K, Aoki Y et al (2012) Novel protein extraction approach using micro-sized chamber for evaluation of proteins eluted from formalin-fixed paraffinembedded tissue sections. Proteome Sci 10:19 10. Shi SR, Taylor CR, Fowler CB et al (2013) Complete solubilization of formalin-fixed, paraffin-embedded tissue may improve proteomic studies. Proteomics Clin Appl 7(3–4): 264–272 11. Xiao Z, Li G, Chen Y et al (2010) Quantitative proteomic analysis of formalin-fixed and paraffin-embedded nasopharyngeal carcinoma using iTRAQ labeling, two-dimensional liquid chromatography, and tandem mass spectrometry. J Histochem Cytochem 58(6):517–527 12. Ono A, Kumai T, Koizumi H et al (2009) Overexpression of heat shock protein 27 in

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squamous cell carcinoma of the uterine cervix: a proteomic analysis using archival formalinfixed, paraffin-embedded tissues. Hum Pathol 40(1):41–49 Jain MR, Liu T, Hu J et al (2008) Quantitative proteomic analysis of formalin fixed paraffin embedded oral HPV lesions from HIV patients. Open Proteomics J 1:40–45 Donadio E, Giusti L, Cetani F et al (2011) Evaluation of formalin-fixed paraffinembedded tissues in the proteomic analysis of parathyroid glands. Proteome Sci 9(1):29 Ostasiewicz P, Zielinska DF, Mann M et al (2010) Proteome, phosphoproteome, and N-glycoproteome are quantitatively preserved in formalin-fixed paraffin-embedded tissue and analyzable by high-resolution mass spectrometry. J Proteome Res 9(7):3688–3700 Azimzadeh O, Scherthan H, Yentrapalli R et al (2012) Label-free protein profiling of formalinfixed paraffin-embedded (FFPE) heart tissue reveals immediate mitochondrial impairment after ionising radiation. J Proteomics 75(8): 2384–2395 Hauck SM, Dietter J, Kramer RL et al (2010) Deciphering membrane-associated molecular processes in target tissue of autoimmune uveitis by label-free quantitative mass spectrometry. Mol Cell Proteomics 9(10):2292–2305 Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227(5259):680–685 Thomas PD, Kejariwal A, Guo N et al (2006) Applications for protein sequence-function evolution data: mRNA/protein expression analysis and coding SNP scoring tools. Nucleic Acids Res 34(Web Server issue):645–650 da Huang W, Sherman BT, Lempicki RA (2009) Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources. Nat Protoc 4(1):44–57 Ashburner M, Ball CA, Blake JA et al (2000) Gene ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat Genet 25(1):25–29 Fowler CB, Cunningham RE, O'Leary TJ et al (2007) 'Tissue surrogates' as a model for archival formalin-fixed paraffin-embedded tissues. Lab Invest 87(8):836–846 Nirmalan NJ, Harnden P, Selby PJ et al (2008) Mining the archival formalin-fixed paraffinembedded tissue proteome: opportunities and challenges. Mol Biosyst 4(7):712–720

Chapter 11 Full-Length Protein Extraction Protocols and Gel-Based Downstream Applications in Formalin-Fixed Tissue Proteomics Alessandro Tanca, Sergio Uzzau, and Maria Filippa Addis Abstract Archival formalin-fixed, paraffin-embedded (FFPE) tissue repositories and their associated clinical information can represent a valuable resource for tissue proteomics. In order to make these tissues available for protein biomarker discovery and validation studies, dedicated sample preparation procedures overcoming the intermolecular cross-links introduced by formalin need to be implemented. This chapter describes a full-length protein extraction protocol optimized for downstream gel-based proteomics applications. Using the procedures detailed here, SDS-PAGE, western immunoblotting, GeLC-MS/MS, 2D-PAGE, and 2D-DIGE can be carried out on FFPE tissues. Technical tips, critical aspects, and drawbacks of the method are presented and discussed. Key words FFPE, Paraffin-embedded, Biobanks, Archival tissues, GeLC-MS/MS, Proteomics, 2D-PAGE, DIGE, Electrophoresis, Immunoblotting, Mass spectrometry

1

Introduction Tissue biopsies are subjected to formalin fixation and paraffin embedding (FFPE) in order to enable their microscopic examination in the context of routine diagnostic procedures. Once the diagnostic process is completed, the residual tissues are stored in long-term repositories. Clearly, these archives possess a huge potential for retrospective tissue proteomics studies, thanks to their matched datasets on the patient clinical records including diagnosis, disease progression, and response to therapy. In addition, when considering the restrictions due to significant ethical issues, logistic complexity, or poor availability of rare disease samples, that are connected with the recovery of sufficient numbers of high-quality, well-characterized fresh tissues, archived FFPE tissues become an especially valuable resource [1, 2].


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Nevertheless, the formalin fixation chemistry introduces stable intermolecular cross-links within the tissue. As a consequence, until less than a decade ago it was practically impossible to access proteomic information of FFPE tissues. In the latest years, and with varying degrees of success, several research groups have worked at the development of methods aimed to solve this problem, leading to the devisal of strategies that fall into two main approaches: digestion of the whole cross-linked protein matrix into peptides, or extraction of full-length proteins by the combined action of physical and chemical factors [3–5]. Typically, direct tryptic digestion strategies generate products that are ideally suitable to liquid chromatography-tandem mass spectrometry (LCMS/MS) analysis, followed by various bioinformatic pipelines for data parsing and interpretation (shotgun proteomics) [6, 7]. Notwithstanding the elevated performances of shotgun proteomics in terms of information depth and data robustness, the ability to obtain full-length proteins can still offer its own advantages, by enabling the application of a wider range of downstream analytical approaches, including separation by gel electrophoresis [8–10]. Each electrophoresis-based method bears its own specific advantages, but all share the added informational value of preserving information on molecular weight or charge of the protein. Among 1D-based approaches, the potential of making FFPE extracts available to SDS-PAGE and western immunoblotting is clearly evident and unquestioned. Despite the outstanding technical progresses made in the proteomics field, western immunoblotting still remains the gold standard for large-scale protein expression validation, due to its technical reliability combined with the relative ease of execution and the low cost of instrumentation and reagents. SDS-PAGE separation of FFPE extracts can also be followed by band cutting, in-gel digestion and MS/MS identification (GeLC-MS/MS), in an alternative to the shotgun proteomics approach. The advantages of GeLC-MS/MS are the ability to process extracts directly without sample cleanup, the direct visualization of the approximate amount and quality of proteins being processed, the maintenance of molecular weight information, as well as the ability to carry out data analysis by using merged information or only information on molecular weight areas of interest [11]. In addition to 1D separations, 2D-PAGE approaches, including the 2D-DIGE technology, can also be implemented on FFPE tissue extracts upon removal of non-compatible extraction reagents. Notwithstanding the tremendous rise and potential of shotgun proteomics approaches, 2D-PAGE is still one of the workhorses of differential proteomics, being accessible, relatively easy to implement, and not strictly dependent from the use of high-performance mass spectrometry instrumentation or data processing capabilities. This widespread technique can provide information on the global patterns of protein expression, enabling the quantitative and

FFPE Tissue Protein Extraction and Analysis

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qualitative study of all proteins in relation to each other without requiring their previous identification, with the added advantage of visualizing proteoforms of different charges or molecular masses [12]. On the other hand, 2D-PAGE has well-known limitations, including poor reproducibility, low sensitivity, and narrow linear dynamic ranges, but these can be overcome by implementation of the 2D-DIGE, that entails the use of fluorescent stains, powerful imaging instrumentation, and ad-hoc experimental design approaches, providing a considerable increase in reproducibility, sensitivity, robustness, and linear dynamic range [13, 14]. The protocol described in this chapter enables extraction of full-length proteins from FFPE tissues building on the “antigen retrieval” concept, described in a seminal work by Shi et al., and elaborated in several variants by different research groups [15, 16]. Here, protein extraction relies on exposure of fixed proteins to combined physical agitation, heat, detergents and reducing agents, in a buffered environment (Fig. 1a). The extracts generated by this protocol are readily suitable to protein quantitation and SDSPAGE separation, followed either by transfer onto nitrocellulose membranes for probing with antibodies, or to in-gel digestion and MS identification. In order to enable 2D-PAGE separation, the ionic detergents used for extraction are exchanged with other reagents suitable to isoelectric focusing of proteins.

Fig. 1 Schematic workflow of the protocol. (a) Protein extraction steps. (b) Downstream gel-based applications

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Once cleared by these interfering substances, proteins can also be subjected to cyanine labeling before separation and undergo the more reliable and robust 2D DIGE workflow (Fig. 1b). The successful implementation of this protocol enables the insertion of FFPE tissue extracts into already established proteomic analysis pipelines. However, it should always be kept in mind that, although extraction and analysis procedures have now reached satisfactory quality levels, FFPE patterns are still not totally comparable to those generated using fresh-frozen tissues. Slight differences do in fact remain concerning high molecular weight and basic proteins. Furthermore, issues such as length of fixation time and tissue cellularity can impact on quality and complexity of the protein pattern, as discussed and detailed previously in several dedicated works [17–19]. In conclusion, when high-quality, fresh tissues are not readily available or practical, the application of the described protocols opens the valuable possibility to exploit the FFPE tissue archive resource for proteomics studies.

2

Materials All protocols require the use of a microcentrifuge and of 1.5 ml microcentrifuge tubes. All reagents may be purchased from SigmaAldrich, if not otherwise specified.

2.1 Tissue Deparaffinization and Rehydration

1. Formalin-fixed paraffin-embedded tissue sections (preferably 5–10 sections, 3–10 μm thick, placed into safe-lock microcentrifuge tubes) (see Notes 1 and 2). 2. Xylene (see Note 3). 3. Ethanol (absolute, 96 and 70 %).

2.2 Protein Extraction and Quantification

1. Extraction buffer: 2 % (w/v) sodium dodecyl sulfate (SDS), 200 mM dithiothreitol (DTT), 20 mM Tris–HCl, pH 8.8 (see Notes 4 and 5). 2. Thermomixer (specifications: temperature up to 100 °C, mixing speed up to 500 rpm; e.g. Thermomixer® Comfort from Eppendorf). 3. Detergent and reducing agent compatible protein quantitation kit (e.g. EZQ™ protein quantitation kit from Life Technologies) (see Note 6).

2.3

SDS-PAGE

1. Laemmli sample buffer: 2 % (w/v) SDS, 10 % (v/v) glycerol, 5 % (v/v) beta-mercaptoethanol, 62.5 mM Tris–HCl, pH 6.8, bromophenol blue (as much as needed). 2. Thermoblock (specifications: temperature up to 100 °C; e.g. Thermomixer® Comfort from Eppendorf).


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3. Precast mini-format polyacrylamide gel (e.g. 4–20 % Mini-PROTEAN® TGX™ Gel from Bio-Rad) (see Note 7). 4. Apparatus for mini-format polyacrylamide gel electrophoresis (e.g. Mini-PROTEAN® Tetra Cell from Bio-Rad). 5. Power supply (e.g. PowerPac™ Basic Power Supply from Bio-Rad). 6. Coomassie-based staining solution SafeStain from Life Technologies).

(e.g.

SimplyBlue™

7. Microwave oven. 2.4

Western Blotting

1. Apparatus for wet electrophoretic protein transfer (e.g. Mini Trans-Blot® Electrophoretic Transfer Cell from Bio-Rad) (see Note 8). 2. Whatman® 3MM cellulose chromatography paper. 3. Nitrocellulose membrane (e.g. Hybond® ECL™ membrane from GE Healthcare) (see Note 9). 4. Transfer buffer: 25 mM Tris, 192 mM glycine, 20 % methanol (see Note 10). 5. Stir bar and magnetic stirrer. 6. Freezer pack. 7. High-current power supply (e.g. PowerPac™ HC HighCurrent Power Supply from Bio-Rad). 8. Ponceau S solution (see Note 11). 9. Bovine Serum Albumin (BSA), lyophilized powder. 10. Phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, pH 7.4. 11. PBS-T solution: 0.05 % Tween-20 in PBS. 12. Blocking solution: PBS-T plus 3 % BSA. 13. Primary antibody solution: primary antibody diluted in PBS-T plus 1 % BSA. 14. Secondary antibody solution: peroxidase-conjugated secondary antibody diluted in PBS-T plus 1 % BSA. 15. Chemiluminescent peroxidase substrate. 16. Imaging system suitable for chemiluminescent signal acquisition (e.g. Molecular Imager® VersaDoc™ MP imaging system from Bio-Rad).

2.5 2-D DIGE and 2-D PAGE

1. 2-D Clean-Up Kit (GE Healthcare). 2. TUC buffer: 7 M urea, 2 M thiourea, 4 % (w/v) 3-[(3Cholamidopropyl)dimethylammonio]-1- propanesulfonate (CHAPS), 10 mM Tris–HCl, pH 8.8. 3. CyDye™ DIGE Fluors (GE Healthcare).

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4. N,N-Dimethylformamide (DMF) anhydrous, >99 % (see Note 12). 5. Immobilized pH gradient (IPG) 24 cm strips (e.g. Immobiline® DryStrip from GE Healthcare) (see Note 13). 6. Carrier ampholytes (e.g. IPG buffer from GE Healthcare). 7. 2-D reducing buffer: 8 M urea, 130 mM DTT, 4 % (w/v) CHAPS, 2 % (v/v) IPG Buffer. 8. Rehydration buffer: 8 M Urea, 13 mM DTT, 4 % (w/v) CHAPS, 1 % (v/v) IPG Buffer. 9. Rehydration tray for IPG strips (e.g. IPG box, along with a reswelling tray, from GE Healthcare) (see Note 14). 10. Isoelectric focusing (IEF) apparatus (e.g. Ettan™ IPGphor 3 from GE Healthcare) with IPG manifold. 11. Paper wicks (two per strip). 12. Twenty-four centimeter gel casting system (e.g. Ettan™ DALTtwelve Gel Caster from GE Healthcare). 13. Gradient former (e.g. Model 495 Gradient former from Bio-Rad). 14. “Light” solution for gradient gel casting: 10 % (v/v) Acrylamide/Bis-acrylamide (29:1) solution, 0.375 M Tris– HCl, pH 8.8, 0.05 % (w/v) ammonium persulfate (APS), 0.025 % (w/v) N,N,N′,N′-Tetramethylethylenediamine (TEMED). 15. “Heavy” solution for gradient gel casting: 18 % (v/v) Acrylamide/Bis-acrylamide (29:1) solution, 0.375 M Tris– HCl, pH 8.8, 10 % (v/v) glycerol, 0.05 % (w/v) APS, 0.025 % (w/v) TEMED. 16. Displacing solution: 50 % (v/v) glycerol, 0.375 M Tris–HCl, pH 8.8, bromophenol blue (as much as needed). 17. Water-saturated n-butanol: 70 % (v/v) n-butanol. 18. Equilibrating buffer: 6 M urea, 2 % (w/v) SDS, 30 % (v/v) glycerol, 50 mM Tris–HCl, pH 8.8. 19. Equilibrating-reducing buffer: equilibrating buffer with 2 % (w/v) DTT. 20. Equilibrating-alkylating buffer: equilibrating buffer with 2.5 % (w/v) iodoacetamide (IAM). 21. Equilibration tray. 22. Agarose solution: 0.4 % (low melting point) agarose, 0.375 M Tris–HCl, pH 8.8, bromophenol blue (as much as needed). 23. Second dimension apparatus (e.g. Ettan™ DALTtwelve system from GE Healthcare). 24. Running buffer: 25 mM Tris, 192 mM glycine, 0.1 % SDS.


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25. Laser scanner suitable for fluorescent sample imaging (e.g. Typhoon™ Trio + laser scanner from GE Healthcare). 26. Software for differential analysis of 2D-DIGE images (e.g. DeCyder™ 2-D Differential Analysis Software, version 7.0 or higher, from GE Healthcare). 27. Fixing solution: 40 % methanol, 10 % acetic acid (see Note 15). 28. Colloidal Coomassie dye (e.g. HPE™ Coomassie® Staining Kit from SERVA). 29. Flatbed scanner suitable for densitometric analysis of 2D-gels (e.g. ImageScanner™ III from GE Healthcare). 2.6 In Gel Trypsin Digestion

1. Clean scalpel. 2. Acetonitrile (ACN). 3. 50 mM ammonium bicarbonate (ABC) (see Note 16). 4. Reducing solution: 10 mM DTT, 50 mM ABC (see Note 5). 5. Alkylating solution: 55 mM IAM, 50 mM ABC. 6. Trypsin, proteomics grade. 7. Air circulation thermostat. 8. 20 % trifluoroacetic acid (TFA). 9. Vacuum centrifuge (e.g. Concentrator plus from Eppendorf). 10. 0.2 % formic acid (FA).

3

Methods

3.1 Tissue Deparaffinization and Rehydration

1. Add 1 ml xylene per each tube containing the FFPE tissue sections. 2. Vortex 10 s and incubate 5 min at RT. 3. Centrifuge 3 min at 16,000 × g at RT. 4. Remove supernatant (see Note 17). 5. Repeat steps 1–4 twice. 6. Add 1 ml 100 % ethanol per tube. 7. Repeat steps 2–4. 8. Add 1 ml 96 % ethanol per tube. 9. Repeat steps 2–4. 10. Add 1 ml 70 % ethanol per tube. 11. Repeat steps 2–4. 12. Spin down, remove all remaining supernatant and let the tube open in the hood for 5 min to dry the pellet (see Note 18).

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3.2 Protein Extraction and Quantification

1. Add the extraction buffer to the tissue pellet (usually 100 μl buffer per 10 mg tissue) and gently mix with the pipette tip (without pipetting or vortexing) (see Note 19). 2. Incubate at 99 °C for 20 min at 500 rpm in a thermomixer. 3. Incubate at 80 °C for 2 h at 500 rpm in a thermomixer (see Note 20). 4. Centrifuge for 10 min at 16,000 × g, collect the supernatant and transfer it to a clean tube (see Note 21). 5. Quantify the protein extract with a detergent and reducing agent compatible quantitation kit and store it at −20 or −80 °C (see Note 6).

3.3 Western Immunoblotting

1. Mix the protein extract(s) (usual load 1–10 μg per well) with an equal volume of Laemmli sample buffer (see Note 22). 2. Incubate the sample(s) for 5 min at 95 °C in a thermoblock, then spin down briefly. 3. Assemble the apparatus for mini-format polyacrylamide gel electrophoresis, add an adequate volume of running buffer and load the sample(s). 4. Connect the gel electrophoresis apparatus to the power supply and run according to the manufacturer’s instructions (approx. 30 min at 200 V for Bio-Rad TGX gels). 5. Equilibrate sponges, 3MM paper sheets, polyacrylamide gel (containing the separated protein sample) and nitrocellulose membrane by soaking them for a few seconds in transfer buffer. 6. Assemble a “sandwich” composed by: sponge, two 3MM paper sheets, polyacrylamide gel, nitrocellulose membrane, two 3MM paper sheets, sponge (from the black cathode to the red anode). 7. Put the sandwich into the transfer apparatus, along with a prefrozen freezer pack and a stir bar. 8. Position the transfer apparatus on a magnetic stirrer and fill it up with transfer buffer. 9. Connect the transfer apparatus to the power supply and run at 250 mA for (at least) 1 h (see Note 23). 10. Stain the membrane with Ponceau staining for a few seconds and destain with deionized water until the background is almost completely white (if necessary, acquire the image with a scanner) (see Note 11). 11. Block the membrane for 1 h with blocking solution (see Note 24). 12. Remove the blocking solution, pour an adequate volume of PBS-T and wash the membrane with rapid agitation for 5 min.


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13. Remove PBS-T, add the primary antibody solution and incubate with gentle agitation for 1 h (after incubation, collect the primary antibody solution if planning to re-use it) (see Note 24). 14. Wash the membrane with an adequate volume of PBS-T with rapid agitation for 5 min. 15. Repeat step 14 twice. 16. Remove PBS-T, add the secondary antibody solution and incubate with gentle agitation for 1 h (after incubation, remove the secondary antibody solution). 17. Repeat step 14 five times. 18. Wash the membrane with an adequate volume of PBS (alone, without Tween-20) with rapid agitation for 5 min. 19. Remove excess PBS from the membrane (without allowing it to dry out completely). 20. Put the membrane on a glass covered by a plastic film, cover it with an adequate volume of chemiluminescent substrate and incubate for 5 min. 21. Remove excess substrate from the membrane (without allowing it to dry out completely) and put the membrane between two tracing paper sheets. 22. Acquire the chemiluminescent signal with a suitable imaging system (see Note 25). 3.4

GeLC-MS/MS

1. Separate protein extract(s) by SDS-PAGE, according to steps 1–4 of Subheading 3.3, except the total sample load (usually 20–30 μg per lane). 2. Stain the gel with a Coomassie-based dye (e.g. three sequential 1 min microwave incubations in deionized water followed by a 45 s microwave incubation with the staining solution if using the SimplyBlue SafeStain staining). 3. Put the stained gel on a clean glass. 4. Fractionate the entire lane(s) in 15–30 slices using a clean scalpel, and transfer each slice into a clean tube (see Note 26). 5. Shrink gel slices by adding ACN (up to completely cover the gel piece) and incubating for 10 min at RT (gel pieces become opaque and stick together). 6. Completely remove ACN, add the reducing solution up to completely cover gel pieces and incubate for 45 min at 56 °C (see Note 5). 7. Chill tubes to RT and remove the remaining solution. 8. Add ACN up to completely cover gel pieces and incubate for 10 min at RT.

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9. Completely remove ACN, add the alkylating solution up to completely cover gel pieces and incubate for 30 min at RT in the dark. 10. Remove the remaining solution, add ACN up to completely cover gel pieces and incubate for 10 min. 11. Completely remove ACN. 12. If gel pieces are not completely destained (blue is still visible) add 50 mM ABC and incubate at RT for 10 min, then shrink with ACN for 10 min (repeat sequentially until gel pieces are destained); otherwise, skip to the following step (see Note 27). 13. Dissolve trypsin in ABC in order to reach a 10 ng/μl working solution. 14. Add 10 μl of trypsin working solution to the dry gel pieces and leave them in an ice bucket or at 4 °C for 60–120 min. 15. Remove excess of trypsin solution, then add ABC up to completely cover gel pieces. 16. Place tubes with gel pieces into an air circulation thermostat and incubate overnight at 37 °C. 17. Chill tubes to RT, spin down and transfer each ABC supernatant to a new tube. 18. Add ACN up to completely cover gel pieces and incubate for 10 min. 19. Collect the ACN supernatant and add it to the tube containing the ABC supernatant belonging to the same gel slice (see Note 28). 20. Acidify by adding into each tube 10 μl of 20 % TFA per 100 μl of peptide solution. 21. Dry the solution in a vacuum centrifuge. 22. Resuspend the pellet in 0.2 % FA and store at −20 or −80 °C until mass spectrometry (MS) analysis. 3.5 Gradient Gel Casting for 2D-DIGE

1. Fill the gel caster by alternating cassettes with separator sheets, then tighten all the screws evenly. 2. Place the gradient former on a magnetic stir plate and add a stir bar to the mixing chamber labeled “light” (both stopcocks and the black lever should be in the closed position). 3. Prepare the “heavy” and “light” acrylamide solutions, except APS and TEMED, and the displacing solution. 4. Immediately prior to pouring, add TEMED and APS to both solutions, mix evenly, then gently pour the appropriate solutions into the gradient chambers (see Note 29). 5. Turn on the stirring bar in the mixing chamber to a steady speed and maintain this same speed throughout casting.


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6. Start casting the gels by opening the stopcock valve to the multi-casting chamber as well as the valve closer to the gradient former (the black lever on the gradient former is still closed at this time) (see Note 30). 7. Allow the light monomer solution to enter the multi-casting chamber until the level of solution in the mixing chamber is equal to the level of the “heavy” solution in the reservoir chamber, then open the black lever on the gradient former and begin mixing the solutions and creating the gradient. 8. When the acrylamide solution in the gradient chamber has almost finished, close the stopcock on the gradient former and add to the reservoir chamber about 200 ml of displacing solution. 9. Open the stopcock on the gradient former and allow the displacing solution to enter the multi-casting chamber until the acrylamide solution has reached the desired level at the top. 10. Close the stopcock on the multi-casting chamber, immediately pipette water-saturated n-butanol onto each gel to overlay, and allow gels to polymerize at RT for at least 12 h (see Note 31). 3.6

2D-DIGE

The following protocol has been optimized using the GE Healthcare materials and apparatus suitable for 2D-DIGE analysis. Alternative reagents and instruments might also be used, but refinements to the protocol might be therefore necessary. 1. Precipitate the protein extract(s) to be analyzed by 2D-DIGE it with the 2-D Clean-Up Kit, following the manufacturer’s instructions (see Note 32). 2. Resuspend the protein pellet with the TUC buffer, for a final concentration of about 5 μg/μl (see Note 33). 3. Add 1.5 volumes of DMF to 1 volume of CyDye stock solution, to make a 400 pmol/μl CyDye working solution. 4. Pipette a volume of protein sample equivalent to 50 μg into a clean tube, then add 1 μl of CyDye working solution, mix by pipetting and leave on ice for 30 min in the dark. 5. Add 1 μl of 10 mM lysine to stop the reaction, mix by pipetting, and leave for 10 min on ice in the dark. 6. Add an equal volume of 2-D reducing buffer to each sample and leave on ice for 10 min (see Note 34). 7. Mix the labeled protein samples that are going to be separated on the same gel. 8. Add an adequate volume of rehydration buffer up to 450 μl in total (for 24 cm strips). 9. Pipette the sample evenly over a slot of the IPG box (see Note 35).

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10. Carefully pull off the cover film from the IPG strip gel and place the strip into the slot, gel-side down, so that the rehydration solution can distribute evenly under the strip. 11. Gently close the lid of the IPGbox and allow the IPG strip gels to rehydrate at RT for 6–24 h (see Note 36). 12. Transfer the strips from the IPGbox to the IPGphor manifold, so that strips are faced up under the cover fluid with the anodic end (+) pointing at the anode of the IPGphor. 13. Add 150 μl deionized water to each paper wick (two per strip), and position the wicks on each end of the IPG strips so that one end of the wick overlaps the end of the IPG strip gel. 14. With the electrode cams in the open position, put the electrode assembly on top of all the wicks (the electrode must be in contact with the wick), then close the lid and start the appropriate IEF program. 15. Just before the end of the IEF program, prepare the equilibration-reduction solution by adding 20 mg DTT per ml of equilibration buffer. 16. Stop the IEF, drain the excess of mineral oil and briefly rinse out the IPG strips with deionized water (see Note 37). 17. Position each IPG strip in a slot of the equilibration tray, add an adequate volume of equilibration-reduction solution up to completely cover the strip and incubate for 15 min with gentle agitation. 18. Turn the pump valve of the Ettan DALTtwelve to “circulate,” then fill the tank to the 7.5 l line with 1× running buffer. 19. On the control unit, adjust the temperature to the desired setting (25 °C) and turn the pump on. 20. Prepare the equilibration-alkylation solution by adding 25 mg IAM per milliliter of equilibration buffer. 21. Remove DTT solution from the IPG strips, add an adequate volume of equilibration-alkylation solution up to completely cover the strip and incubate for 15 min with gentle agitation. 22. Carefully unload the cassettes containing the polymerized gels from the gel caster and rinse the top surface and the glass plates with deionized water to remove residual n-butanol and acrylamide. 23. Using forceps, remove the equilibrated IPG strips from the equilibration solution and rinse them with fresh SDS running buffer. 24. Slowly pipette agarose solution on the top of each gel, then position the strip centered on the glass plate, and push against the plastic backing of the IPG strip (with forceps or a plastic support) to slide the strip between the two glass plates until it is into contact with the surface of the slab gel.


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25. Carefully slide the gels into the tank and fill the upper buffer chamber with 2.5 l of 2× running buffer (up to the min/max fluid line) (see Note 38). 26. Close the lid, set the running conditions according to the desired run length (for instance, 1 W per gel overnight) and start the run. 27. When the run has finished, unload the gel cassettes from the electrophoresis chamber, rinse them with deionized water and proceed with image acquisition using Typhoon laser scanner. 28. Upload the image files into the DeCyder software and carry out differential analysis according to the software manual. 3.7 Preparation of a Preparative Gel, Excision, and Digestion of Differential Spots

1. Collect a volume of the protein sample obtained in step 2, Subheading 3.6, corresponding to at least 300 μg of samples (preferably 500–700 μg). 2. Perform 2D-PAGE analysis according to steps 8–26 of Subheading 3.6. 3. When the run has finished, unload the cassette from the electrophoresis chamber, open it and carefully put the gel into a clean container. 4. Fix the gel with fixing solution for 30–60 min, stain overnight with a colloidal Coomassie dye, destain with deionized water and proceed with image acquisition using a flatbed scanner (see Note 39). 5. Excise from the preparative gel all the differential spots identified upon DeCyder analysis. 6. Shrink gel slices by adding ACN (up to completely cover the gel piece) and incubating for 10 min at RT. 7. Perform in gel digestion and peptide recovery according to steps 11–22 of Subheading 3.4.

4

Notes 1. FFPE tissue blocks should be carefully selected and evaluated by an expert pathologist in order to assure the presence of a minimum percentage of cells of the desired tissue type (for instance, >90 % of neoplastic tissue), and consecutive sections should be obtained for proteomic analysis. In general, five 5-μm-thick tissue sections should provide sufficient material for a gel-based proteomic analysis. Furthermore, especially for precious tumor samples, an additional step of manual dissection or laser microdissection can be favorably added to enrich in cancer cells and keep the amount of adjacent connective and normal tissue to the minimum.

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2. Safe-lock tubes should be used, since a loose locking may cause the tube to “uncork” during the high temperature incubation steps, with a consequent loss of biological material. 3. Xylene may cause harmful effects by eye and skin contact as well as by inhalation, ingestion or aspiration, spanning from minor irritation to headache, nausea and vomiting. It has therefore to be used with care, wearing gloves and under a chemical hood. As an alternative for tissue deparaffinization, a mineral oil mixture (heated at 95 °C) can be used with similar results. 4. Several different extraction buffers have been described in the literature as suitable for extraction of full-length proteins from FFPE tissue samples. SDS and DTT are both widely recognized as essential to ensure a proper extraction and solubilization of proteins, although their concentration may vary (e.g. 4 % SDS is well documented). The extraction buffer can also comprise additional reagents, such as octyl glucoside, glycerol and polyethylene glycol [4]. 5. All solutions containing DTT should be prepared fresh shortly before use. Alternatively, stock solutions without DTT may be prepared, and an appropriate amount of DTT powder may be added just prior to use. 6. The use of a protein quantitation method being compatible with detergents (SDS) and reducing agents (DTT) is mandatory. The fluorescence-, membrane-based EZQ™ system is the most used in our lab with FFPE protein extracts for three reasons: (a) it requires a very low volume of sample for quantification (1 μl); (b) it provides a satisfactory level of linearity and sensitivity even for solutions with low protein concentrations; (c) the washing steps enable an efficient removal of most detergent and reducing agent. These characteristics fit well with the very low size of many FFPE tissue samples, and therefore with the low amount and/or concentration of the corresponding protein extracts. Anyway, comparable or even better quantification performance can be achieved using alternative detergent and reductant compatible methods. 7. It has been demonstrated that FFPE protein extracts contain a lower amount of high molecular weight (MW) proteins compared to fresh-frozen tissues; a general depletion in the upper part of 1D- or 2D-PAGE pattern can be therefore observed in most cases. According to this, in order to enhance resolution in the medium–low MW part of the pattern, an acrylamide percentage higher than that used for other samples can be conveniently chosen for gel-based separation (e.g. 4–20 % for precast gradient gels and 10–18 % for home-made, large format gradient gels; 14 % fixed-percentage gels can also be used). 8. Alternatively, semi-dry or dry protein transfer systems could be used with similar results.


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9. Alternatively, a PVDF membrane can be employed. In this case, preliminary activation of PVDF membrane with methanol is needed. 10. Up to 0.1 % SDS may be added to the transfer buffer to maximize transfer of hydrophobic proteins. 11. Membrane staining with Ponceau is aimed at: (a) verifying if bubbles have formed which hamper a complete transfer of the proteins from certain regions of the gel; (b) checking if the total load of the different samples is actually comparable; (c) acquire an image of the total protein pattern which can be overlapped to the final antibody signal as a control. 12. DMF can have carcinogen and teratogen effects, and thus needs to be used with great care, wearing gloves and under a chemical hood. It is also very important that a high grade, anhydrous DMF is used to ensure an efficient labeling. Use a DMF bottle less than 3 months old from day of opening and prevent it from being exposed to air or water. 13. Using a large strip format (24 cm) is advised to reach a high resolution and a proper separation of protein spots. 14. The IPGbox kit allows the passive rehydration to be performed in the dark and, more importantly, without the need for covering the strips with mineral oil. 15. Some research groups have replaced methanol (more toxic) with ethanol (safer, but slightly less efficient). 16. The ABC solution should be filtered before use and stored at 4 °C to avoid contaminations. 17. Take particular care when removing supernatant in order not to throw away tissue portions. If extracting proteins from filamentous or colloidal tissues, increasing centrifugation time may help pellet the tissue more efficiently. 18. Tissue pellets may also be vacuum dried, in order to maximize absorption and penetration of the extraction buffer in the next step. However, special attention to be paid to the vacuum centrifugation time to avoid excessive drying (or even burning) which would make it difficult or impossible to resuspend the tissue in the extraction buffer. 19. Usually gently mixing with a tip (a movement approximately like stirring coffee with a spoon) is enough to adequately ensure contact between buffer solution and tissue fragments. Strictly avoid vortexing or hard pipetting, because of the presence of SDS in the buffer. Alternatively, a tissue disruption step (e.g. performed using a steel bead and a mechanical homogenizer) may allow a deeper and more even penetration of the extraction buffer within the tissue. 20. At this stage, an overnight storage at −80 °C may help improve lysis and protein release from the tissue.

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21. If a clear separation between the residual pellet and the protein containing supernatant does not occur, collect the greatest possible volume of supernatant without touching the tissue, then centrifuge again the remaining “wet” pellet for at least 15 min at 16,000 × g to “squeeze” it. Finally, collect the new supernatant and merge it with the one obtained previously. 22. The protein load should be settled depending on the following factors: (a) concentration/abundance of the target antigen within the protein mixture; (b) specificity of the antibody; (c) sensitivity of the chemiluminescent substrate and of the signal detection system. For a medium–low abundance antigen, a monoclonal antibody, the Sigma chemiluminescent peroxidase substrate and the VersaDoc imager, 1 μg of total protein should be enough. 23. When interested in high MW proteins, the transfer time may be extended to 1.5–2 h. 24. Incubations may be extended up to overnight based on sample or antibody characteristics. For overnight incubations, preincubate the membrane at RT for approx. 30 min, then keep the membrane at 4 °C during the night, and finally equilibrate the membrane the day after at RT for other 30 min. 25. If using the VersaDoc imaging system, a 5 min exposure is usually enough to detect a visible signal. 26. The number of slices in which the lane is cut (i.e. the degree of sample fractionation) is usually proportional to the number of protein identifications achieved through MS analysis. Alternatively, the number of protein identifications can be also increased by increasing the length of the LC separation prior to MS (a detailed description of the LC-MS/MS analysis methods is outside the scope of this chapter). 27. Alternatively, samples can be destained using a solution containing 40 % (v/v) ethanol and 75 mM ABC and/or incubating the samples at 56 °C and 500 rpm in a thermomixer until the blue dye is no longer visible. 28. If the gel pieces are not completely shrunk and opaque after collecting ACN, add fresh ACN up to cover the gel pieces, incubate for 10 min, collect the new ACN supernatant and add it to the supernatant mixture belonging to the same gel slice. 29. In other words, the solution with the lower acrylamide concentration is to be poured in the mixing chamber labeled “light,” and the solution with the higher acrylamide concentration in the reservoir chamber labeled “heavy.” Note that the level of the solutions in the two chambers will NOT be equal at this time.


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30. Do not allow any air bubbles to enter the plates when casting gradient gels, since bubbles hamper a correct formation of the gradient. 31. The multi-casting chamber should be stored at RT possibly overnight to ensure the complete acrylamide polymerization. In fact, acrylamide polymerization continues in a “silent” way even after gels start to visually appear as polymerized. Afterwards, unused gels can be stored between wet papers at 4 °C for up to 2 weeks. 32. Protein precipitation is performed to remove SDS and make protein solution suitable for IEF separation. Alternatively, detergent removal spin columns, dialysis membranes, or MW-cut-off spin filters can be used to eliminate SDS from the protein extract. 33. At this stage, sample pH should be between 8 and 9 to ensure maximum labeling efficiency. 34. After this step, cyanine-labeled samples can be stored for at least 3 months at −80 °C in the dark. 35. Ensure that no bubbles are trapped within the strips. 36. Overnight rehydration is recommended to allow high MW proteins to enter the gel. 37. Focused strips can be stored at −80 °C for a few weeks. 38. Lubricate glasses with running buffer to prevent the rubber tubing from sticking to the cassettes. 39. Alternatively, gels can be stained with fluorescent dyes (e.g. Sypro Ruby) and their image can be acquired using an imager suitable for fluorescent signal detection. References 1. Klopfleisch R, Weiss AT, Gruber AD (2011) Excavation of a buried treasure – DNA, mRNA, miRNA and protein analysis in formalin fixed, paraffin embedded tissues. Histol Histopathol 26(6):797–810 2. Fowler CB, O'Leary TJ, Mason JT (2013) Toward improving the proteomic analysis of formalin-fixed, paraffin-embedded tissue. Expert Rev Proteomics 10(4):389–400 3. Blonder J, Veenstra TD (2009) Clinical proteomic applications of formalin-fixed paraffinembedded tissues. Clin Lab Med 29(1): 101–113 4. Tanca A, Pagnozzi D, Addis MF (2012) Setting proteins free: progresses and achievements in proteomics of formalin-fixed, paraffin-embedded tissues. Proteomics Clin Appl 6(1–2):7–21

5. Maes E, Broeckx V, Mertens I, Sagaert X, Prenen H, Landuyt B, Schoofs L (2013) Analysis of the formalin-fixed paraffinembedded tissue proteome: pitfalls, challenges, and future prospectives. Amino Acids 45(2):205–218 6. Stewart NA, Veenstra TD (2008) Sample preparation for mass spectrometry analysis of formalin-fixed paraffin-embedded tissue: proteomic analysis of formalin-fixed tissue. Methods Mol Biol 425:131–138 7. Krizman DB, Burrows J (2013) Use of formalin-fixed, paraffin-embedded tissue for proteomic biomarker discovery. Methods Mol Biol 1002:85–92 8. Addis MF, Tanca A, Pagnozzi D, Crobu S, Fanciulli G, Cossu-Rocca P, Uzzau S (2009) Generation of high-quality protein extracts

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9.

10.

11.

12. 13.

14.

Alessandro Tanca et al. from formalin-fixed, paraffin-embedded tissues. Proteomics 9(15):3815–3823 Addis MF, Tanca A, Pagnozzi D, Rocca S, Uzzau S (2009) 2-D PAGE and MS analysis of proteins from formalin-fixed, paraffin-embedded tissues. Proteomics 9(18):4329–4339 Tanca A, Pagnozzi D, Falchi G, Tonelli R, Rocca S, Roggio T, Uzzau S, Addis MF (2011) Application of 2-D DIGE to formalin-fixed, paraffin-embedded tissues. Proteomics 11(5): 1005–1011 Tanca A, Pagnozzi D, Burrai GP, Polinas M, Uzzau S, Antuofermo E, Addis MF (2012) Comparability of differential proteomics data generated from paired archival fresh-frozen and formalin-fixed samples by GeLC-MS/MS and spectral counting. J Proteomics 77:561–576 Chevalier F (2010) Highlights on the capacities of “Gel-based” proteomics. Proteome Sci 8:23 Viswanathan S, Unlu M, Minden JS (2006) Two-dimensional difference gel electrophoresis. Nat Protoc 1(3):1351–1358 Tanca A, Pisanu S, Biosa G, Pagnozzi D, Antuofermo E, Burrai GP, Canzonieri V, Cossu-Rocca P, De Re V, Eccher A, Fanciulli G, Rocca S, Uzzau S, Addis MF (2013)

15.

16.

17.

18.

19.

Application of 2D-DIGE to formalin-fixed diseased tissue samples from hospital repositories: results from four case studies. Proteomics Clin Appl 7(3–4):252–263 Shi SR, Taylor CR, Fowler CB, Mason JT (2013) Complete solubilization of formalinfixed, paraffin-embedded tissue may improve proteomic studies. Proteomics Clin Appl 7(3–4):264–272 Shi S-R, Shi Y, Taylor CR (2011) Antigen retrieval immunohistochemistry. J Histochem Cytochem 59(1):13–32 Tanca A, Pagnozzi D, Falchi G, Biosa G, Rocca S, Foddai G, Uzzau S, Addis MF (2011) Impact of fixation time on GeLC-MS/MS proteomic profiling of formalin-fixed, paraffin-embedded tissues. J Proteomics 74(7):1015–1021 Thompson SM, Craven RA, Nirmalan NJ, Harnden P, Selby PJ, Banks RE (2013) Impact of pre-analytical factors on the proteomic analysis of formalin-fixed paraffin-embedded tissue. Proteomics Clin Appl 7(3–4):241–251 Magdeldin S, Yamamoto T (2012) Toward deciphering proteomes of formalin-fixed paraffin-embedded (FFPE) tissues. Proteomics 12(7):1045–1058

Chapter 12 Enrichment of Low-Abundant Protein Targets by Immunoprecipitation Upstream of Mass Spectrometry Barbara Kaboord, Suzanne Smith, Bhavin Patel, and Scott Meier Abstract Immunoprecipitation (IP) is commonly used upstream of mass spectrometry (MS) as an enrichment tool for low-abundant protein targets. However, several aspects of the classical IP procedure such as nonspecific protein binding to the isolation matrix, detergents or high salt concentrations in wash and elution buffers, and antibody chain contamination in elution fractions render it incompatible with downstream mass spectrometry analysis. Here, we discuss two IP workflows that are designed to minimize or eliminate these contaminants: the first employs biotinylated antibodies and streptavidin magnetic beads while the second method utilizes a traditional antibody that is oriented and cross-linked to Protein AG magnetic beads. Both modified magnetic supports have low background binding and both antibody immobilization strategies significantly reduce or eliminate antibody heavy and light chain contamination in the eluent, minimizing potential ion suppression effects and thereby maximizing detection of target antigens and interacting proteins. Key words Immunoprecipitation, Co-immunoprecipitation, Mass spectrometry, Protein–protein interactions, Biotin, Streptavidin, Magnetic beads, Protein AG, In-solution digestion

1

Introduction Immunoprecipitation is widely used to enrich specific protein targets from complex samples for the purpose of evaluating differential protein expression [1–4], probing for post-translational modifications [5–12], or identifying interacting proteins [13–16]. An antibody is used to bind an antigen in cell lysates, serum, or tissue homogenates and the resulting immune complexes are captured on a solid phase support such as a chromatography resin, magnetic bead, plastic plate, or membrane. Mass spectrometry is increasingly becoming the detection methodology of choice for IPs as the technology has the capability to be used for discovery experiments to identify protein interactors [14, 16–19], semi-quantitatively in conjunction with SILAC or TMT-labeling [20–25], or quantitatively in selective reaction monitoring (SRM) assays [26–30] (Fig. 1).


135

136

Barbara Kaboord et al.

Sample - Cell Culture - Plasma/Serum - Tissue

SDS-PAGE Western blot

Immunoprecipitation

In-Gel/In-Solution Digestion - Reduction/Alkylation - Digestion - Peptide cleanup AQUA Peptide Spiking

Discovery MS (LC-MS/MS) Protein/PTMs Identification, Peptide Selection

Targeted MS (LC-SRM/MS) Absolute Quantitation

Fig. 1 Experimental workflow for IP to MS applications. Low-abundant proteins in a complex sample matrix typically need to be enriched in order to detect or quantify them. Protein targets are immunoprecipitated, processed by in-gel or in-solution digestion, and analyzed by LC-MS/MS to identify proteins in the IP/Co-IP eluent, probe for post-translational modifications, or select candidate peptides. Furthermore, heavy isotope-labeled peptide standards can be added to the sample for targeted LC-SRM/MS for absolute quantitation. Gel electrophoresis followed by silver stain or Western blot is still typically used as a validation component to this workflow

While the best antibody for IP enrichment is highly specific for its target antigen, other abundant proteins from the sample milieu can potentially bind nonspecifically to the antibody, the antigen, or the solid support to which the antibody is immobilized. Buffers containing salt and/or detergent are typically used to eliminate these nonspecific protein binders, however, low-affinity and transient interactions are at risk of being washed away during those steps. In addition, high salt and detergent are not compatible with downstream mass spectrometry analysis. The use of protein crosslinking reagents is a viable strategy to preserve low-affinity complexes [17, 19, 31] during high stringency washes but creates an additional level of complexity during the peptide/protein identification process and still results in high salt or detergent carry-over in the final sample. In addition to abundant protein contaminants from the original sample, antibody chains leach off the supports during the IP elution step and become significant contaminants as well. The presence of large quantities of any type of contaminating protein can suppress ionization of the low-abundant target of interest, preventing its detection in the mass spectrometer. The choice of IP solid support, the antibody attachment strategy, and the IP protocol itself are critical areas to optimize in order to minimize nonspecific protein binding and maximize antigen capture (Fig. 2).

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137

Fig. 2 Evaluation of resin types and IP strategies using an EGFR immunoprecipitation model system. Direct IP (antibody pre-immobilized on the solid support followed by incubation with sample) was compared for three supports (Polyacrylamide (P), Agarose (A) and Magnetic (M)) and four different chemistries of directly coupled antibody. Additionally, biotinylated antibody and two immobilized streptavidin resins were used in an Indirect IP method (antibody + sample preincubation before binding to solid support). (a) Capture efficiency was determined by Western blot. (b) EGFR sequence coverage and background proteins were determined by LC-MS/MS after IP elutions and trypsin digestion. IP using streptavidin magnetic beads resulted in fewer background proteins identified and higher EGFR sequence coverage

Presented here are two IP strategies that minimize both the amount of nonspecific protein binding to the immunoaffinity matrix and the amount of antibody that leaches into the eluted target. The first involves use of a biotinylated antibody and streptavidin magnetic beads. The biotin-labeling process results in multiple biotin modifications per antibody molecule, thus providing multiple contacts or binding points to the streptavidin beads. Because the biotin–streptavidin interaction is extremely high affinity (KD = 10−15 M) and highly resistant to acid elution, the quantity of antibody chains found in the eluent is significantly less than a classical IP with Protein A or G (Fig. 3). Tactically, the use of biotinylated antibodies in IPs is advantageous since biotin labeling is compatible with any antibody species and subtype and utilizes a common streptavidin support matrix. This simplifies the overall workflow and makes the biotin–streptavidin IP approach amenable to multiplexing (e.g. immunoprecipitating and quantifying multiple targets within a cellular signaling pathway). In addition, the biotin/streptavidin approach is preferred with clinical samples such as serum or other biological fluid which normally contains endogenous antibodies that would interfere with binding to the classical Protein AG immunoprecipitation support.


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a

AG + - + - Direct - + - + Indirect

Heavy chain

PP2A Light chain

b Streptavidin Magnec Beads MS results

% Sequence Coverage

Protein AG Magnec Beads

IP/Co-IP targets

PP2A, catalytic subunit (PPP2CA and PPP2CB) PP2A, regulatory subunit (PPP2R1A and PPP2R1B) PP4, catalytic subunit (PPP4C)

Indirect IP

Indirect IP

Direct IP No Crosslink

Direct IP With Crosslink

57%

69%

58%

42%

14%

11%

5%

11%

42%

42%

35%

21%

Fig. 3 Comparison of different IP methods for MS application. (a) Antibody heavy and light chain contamination is significantly reduced in the Protein AG cross-link and the biotinylated antibody/streptavidin methods compared to the classical Protein AG IP methodology. (b) Indirect and direct IP methods were used to evaluate enrichment of PP2A, catalytic subunit (PPP2CA/PPP2CB) from 0.5 mg A431 cell lysate using Streptavidin magnetic beads and Protein AG magnetic beads. IP elutions were processed by in-solution digestion method and analyzed by LC-MS/MS to assess sequence coverage and identify isoform-specific peptides. MS analysis of IP methods using streptavidin and Protein AG magnetic bead supports allowed for identification of two PP2A catalytic subunits (PPP2CA and B), two PP2A regulatory subunits (PPP2R1A and B) and PP4 catalytic subunit (PPP4C). Protein AG cross-link IP method resulted in slight decrease in PPP2CA recovery as indicated by Western blot and by the percent sequence coverage obtained in the mass spectrometry analysis

The second IP strategy is an oriented, cross-linked affinity method [25, 32–34] that uses Protein AG magnetic beads to capture the antibody similar to a classical IP however, before the antigen is introduced, the antibody is covalently cross-linked to the Protein AG using a homobifunctional cross-linker such as DSS. This cross-linking virtually eliminates the presence of IgG heavy and light chain contamination in the eluent (Fig. 3). The oriented, cross-linked approach is a good option if the antibody contains a carrier protein as the binding to Protein AG serves as a purification step before the cross-linking event. Because Protein

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Table 1 Detection and quantification of low-abundant targets by MS Detected by LC-MS/MS

Quantified by LC-SRM/MS

Target

Cell line

Neat

Enriched-IP

Neat

Enriched-IP

EGFR

A431 HEK293

+ −

+ +

+ −

+ +

AKT1

A431 HEK293

− −

+ +

− −

+ +

AKT2

A431 HEK293

− −

+ +

− −

+ +

AKT3

A431 HEK293

− −

+ +

N/A N/A

N/A N/A

PTEN

A431 HEK293

− −

+ +

− −

+ +

PIK3CA

A431

−

+

−

+

PIK3R2

A431

−

+

−

+

KRAS

A431

−

+

N/A

N/A

STAT1

A431

−

+

N/A

N/A

PPP2AC

A431

−

+

N/A

N/A

N/A not available

AG combines the binding characteristics of both Protein A and Protein G, this IP approach also works with a variety of antibody species and subtypes. Classical IP without cross-linking of antibody to Protein AG can always be performed if the presence of antibody chains in the eluent is not a concern (e.g. if performing targeted detection such as SRM assays). IP enrichment is essential to identifying low abundance targets in a complex biological milieu. Many cell signaling proteins and biomarkers are low copy number and are not detected in crude lysates or biological fluids due to the presence of a multitude of other proteins competing for ionization even after LC fractionation. However, with upstream IP enrichment, low-abundant signaling proteins such as EGFR, AKT1, AKT2, PTEN, PIK3CA, PIK3R2, KRAS, and STAT1 can be detected and successfully identified (Table 1). While other biochemical methods of affinity purification upstream of mass spectrometry (AP-MS) exist [35, 36], immunoaffinity purification is used most often for specific enrichment of endogenous protein(s) and is the focus of the protocols described in this chapter.

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2


Materials 1. Pierce Streptavidin Magnetic Beads (Thermo Fisher Scientific, Rockford, IL). Supplied as 10 mg/ml; binding capacity ~55 μg biotinylated rabbit IgG per mg of beads. 2. N-hydroxysuccinimide-PEG4-biotin (NHS-PEG4-biotin), supplied as No-Weigh™ Format (8 × 2 mg in microtubes; MW = 589) (Thermo Fisher Scientific): To make a 20 mM solution, add 170 μl DMF to one 2 mg reagent tube by puncturing the foil seal with the pipette tip. Pipette up and down to mix. Use immediately (see Note 1). 3. Phosphate-buffered saline (PBS): 0.1 M sodium phosphate, 0.15 M NaCl, pH 7.5. 4. Zeba desalting column (0.5 ml; 7K cut-off) (Thermo Fisher Scientific). 5. IP Lysis Buffer (Thermo Fisher Scientific): 25 mM Tris–HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 % NP-40 and 5 % glycerol. 6. Binding/Wash Buffer: 25 mM ammonium bicarbonate, pH 8.0. 7. LC-MS grade water. 8. Elution Buffer: 0.5 % formic acid (LC-MS grade), 30 % acetonitrile (LC-MS grade). 9. Pierce Protein AG Magnetic Beads (Thermo Fisher Scientific). Supplied as 10 mg/ml; binding capacity 55–85 μg rabbit IgG per mg magnetic particles. 10. Disuccinimidyl suberate solution (DSS) (Thermo Fisher Scientific): Dissolve 2 mg DSS in 150 μl DMSO or DMF immediately before use. Once reconstituted, the DSS must be used immediately. DSS is moisture sensitive and must be stored desiccated. 11. Modified PBS: PBS containing 0.05 % NP-40 (see Note 2). 12. 0.1 M glycine-HCl, pH 2.0. 13. Denaturation buffer: 6 M Urea, 50 mM Tris–HCl, pH 8.0 (see Note 3). Add 0.36 g urea (sequencing grade) in 0.5 ml 50 mM Tris–HCl, pH 8.0, to dissolve then bring volume to 1 ml. Do not warm up to dissolve. Prepare fresh for each use. 14. 0.5 M Bond-Breaker TCEP Solution (Thermo Fisher Scientific): Prepare a 1:50 working dilution by adding 10 μl 0.5 M TCEP to 490 μl 50 mM Tris–HCl, pH 8.0. 15. 0.5 M Iodoacetimide: Dissolve 37 mg iodoacetamide in 400 μl MS-grade water. Make fresh and cover with foil to protect from light.

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16. Pierce Trypsin Protease (MS grade): Dissolve 20 μg trypsin in 200 μl 50 mM acetic acid to prepare a 0.1 μg/μl stock trypsin solution. Store aliquots at −80 °C. Prepare (just before use) 10 ng/μl trypsin working solution by adding 10 μl 0.1 μg/μl trypsin stock to 90 μl 50 mM Tris–HCl, pH 8.0. 17. Pierce C18 Spin Tips (Thermo Fisher Scientific). Each tip binds up to 10 μg (20 μl) of total peptide. 18. Trifluoroacetic acid (TFA) (LC-MS grade). 19. Low protein-binding microcentrifuge tubes (Axygen or Eppendorf). 20. Magnetic stand (e.g. DynaMag™-2 Magnet from Life Technologies). 21. Thermomixer. 22. Speed vac. 23. Thermo Scientific™ Orbitrap XL™ mass spectrometer or Thermo Scientific™ Orbitrap™ Fusion™ Tribrid™ mass spectrometer equipped with Thermo Scientific™ Proteome Discoverer™ 1.4 software, Scaffold™ 4.0 (Proteome Software), and SEQUEST®HT search engine (University of Washington).

3

Methods

3.1 Biotinylation of Antibodies and MagneticStreptavidin Immunoprecipitation

The most common approach to biotin-label antibodies is to use N-hydroxysucciminimide (NHS) esters. At pH 7–9, NHS-biotin esters will react specifically with the primary amines in lysine side chains and the N-terminus, creating a stable amide bond. However, the number of biotins incorporated per antibody should be controlled to prevent inactivation of primary amines that may be required for antigen recognition and/or aggregation and precipitation due to over-biotinylation. Utilizing biotinylation reagents with polyethylene glycol (PEG) spacer arms imparts greater water solubility to biotin-labeled antibodies, helping to prevent aggregation. Using the appropriate molar excess of NHS ester over antibody helps titrate the number of biotin labels introduced. In the following protocol, a 40-fold molar excess of biotin was found to result in approximately 2–12 biotin moieties per antibody, a quantity that was found to maximize binding of an antibody on streptavidin beads, preserve antibody function, and minimize leaching of antibody into the eluent due to multiple contact points with the support. 1. Calculate the millimoles of NHS-PEG4-Biotin to add to the labeling reaction to give a 40-fold molar excess over antibody: mg antibody 

mmol antibody 40 mmol biotin   mmol biotin. mg antibody mmol antibody

142


2. Calculate the microliters of 20 mM NHS-PEG4-Biotin to add to each labeling reaction: mmol biotin 

589 mg 170 l   l biotin solution. mmol biotin 2 mg

3. Mix 50–200 μg antibody in PBS (see Note 4) with a 40-fold molar excess of NHS-PEG4-biotin in a total reaction volume of 100 μl. Pipette gently to mix. 4. Incubate the biotin labeling reaction at room temperature for 30 min. Remove excess non-reacted and hydrolyzed biotin reagent by desalting spin column. 5. Centrifuge a 0.5 ml desalting spin column at 1,500 × g for 1 min to remove the storage solution. 6. Equilibrate the desalting column by washing three times with 300 μl PBS. 7. Place desalting column in a fresh collection tube and add the reaction gently to the top of the resin bed. 8. Centrifuge at 1,500 × g for 2 min. Biotinylated antibody is present in the flow-through and is now in PBS (see Note 5). 9. Prepare cell or tissue lysates in IP Lysis Buffer to a final concentration of 1 mg/ml (see Note 6). Sample size per IP reaction is typically 500 μl (500 μg). For serum or plasma samples, dilute 20 μl (~1 mg) of serum or plasma in 480 μl IP Lysis Buffer. 10. For optimal results, pre-clear 500 μl of the lysate sample by incubating with 25 μl of streptavidin magnetic beads for 1 h at room temperature (optional) (see Note 7). 11. Prepare the immune complex by incubating above lysate with 5 μg of antibody overnight at 4 °C with continuous end-overend rotation (see Note 8). 12. Dispense 25 μl streptavidin magnetic beads (for each IP reaction) into a low-binding microcentrifuge tube, collect the beads on a magnetic stand for 1 min and remove the storage solution. Wash the beads twice with IP Lysis Buffer, mixing gently each time to fully suspend the beads. Collect the beads between each wash by placing the tubes on a magnetic stand for 1 min. Discard the supernatants (wash volumes). 13. Add the immune complex prepared in step 11 to the washed streptavidin-magnetic beads and incubate at room temperature for 1 h with end-over-end rotation. 14. Collect the beads on a magnetic stand for 1 min. 15. Gently remove the supernatant (flow-through or non-bound) and save for analysis if desired.

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16. Wash the beads with 3 × 500 μl 25 mM Ammonium bicarbonate, pH 8, by gently vortexing the beads to resuspend during each wash step (see Note 9). 17. Wash the beads with 2 × 500 μl of LC-MS grade water, by gently vortexing the beads to resuspend during each wash step (see Note 10). 18. Elute bound antigen by adding 100 μl elution buffer (0.5 % formic acid, 30 % acetonitrile), and incubating for 5 min at room temperature with periodic gentle vortexing (see Note 11). 19. Collect the beads on a magnetic stand and transfer the elution to a fresh microcentrifuge tube. 20. Dry the elution in a speed vac. 21. Proceed to mass spec sample prep (Subheading 3.3) and/or resuspend in SDS-PAGE sample buffer for Western blot analysis/validation. 3.2 Cross-linked IP Method

An alternative strategy to minimize antibody leaching would be to physically cross-link the antibody to the affinity support either directly to a chemically-activated support or to an immobilized Protein AG bead. The cross-linked Protein AG approach results in an oriented antibody to maximize antigen capture and is the preferred strategy when the antibody contains a carrier protein such as BSA or gelatin. A common homobifunctional cross-linker such as disuccinimidyl suberate (DSS) reacts with amine groups on the antibody and those on the Protein AG support. As with the use of NHS-biotin esters, DSS must be used at the appropriate concentration to minimize inactivation of essential amines in the antigen recognition site that would lead to loss of antibody function. Small decreases in antigen binding are commonly observed with the cross-linked IP method, but this loss of function may not be fully recognized in practice as the immobilized antibody beads are generally still in excess over the target antigen [32, 33] (see Note 12). 1. Vortex the bottle of Pierce Protein AG Magnetic Beads to obtain a homogeneous suspension. Dispense 25 μl of magnetic bead suspension into a low-binding microcentrifuge tube (see Note 13). Place tube on a magnetic stand for 1 min to collect the beads. Remove and discard the storage solution. 2. Wash the beads twice with 500 μl Modified PBS, mixing gently each time to fully suspend the beads. Collect the beads between each wash by placing the tubes on a magnetic stand for 1 min. Discard the supernatants (wash volumes). 3. Dilute antibody in Modified PBS to a final concentration of 5 μg per 100 μl (see Note 14).

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4. Add 100 μl of diluted antibody solution to the washed magnetic beads and incubate at room temperature for 15 min, gently vortexing the beads every 5–10 min during incubation to ensure that the beads stay in suspension. 5. Collect the beads with a magnetic stand. Remove and discard the supernatant. 6. Wash the bound antibody-bead complexes twice with 300 μl Modified PBS. For each wash, thoroughly mix to suspend the beads, collect on a magnetic stand for 1 min, then remove and discard each wash volume. 7. Resuspend the beads in 46 μl PBS. Add 4 μl of 0.25 mM DSS in DMSO or DMF (20 μM final concentration; 10X molar excess DSS over immobilized Protein AG) (see Note 15). 8. Incubate the cross-linker with antibody-Protein AG beads for 30 min at room temperature, vortexing gently every 10–15 min during the incubation to ensure an even bead suspension. 9. Collect the beads with a magnetic stand. Remove the supernatant. 10. Quench the cross-linking reaction and remove non-crosslinked antibody by adding 100 μl 0.1 M glycine buffer, pH 2 to the beads and gently mixing for 5 min at room temperature (see Note 16). Collect the beads with a magnetic stand and remove the supernatant. 11. Perform two additional 100 μl washes with glycine buffer. 12. Equilibrate the cross-linked beads by washing twice with 200 μl cold IP Lysis/Wash Buffer each wash. 13. Prepare cell or tissue lysates in IP Lysis Buffer to a final concentration of 1 mg/ml (see Note 6). Sample size per IP reaction is typically 500 μl (500 μg). For serum or plasma samples, dilute 20 μl (~1 mg) of serum or plasma in 480 μl IP Lysis Buffer. 14. Add 500 μl lysate to the cross-linked magnetic beads and incubate for 1 h at room temperature with end-over-end mixing. Alternatively, vortex the beads every 10–15 min during the incubation to ensure that the beads stay in suspension. 15. Collect the beads on a magnetic stand for 1 min. 16. Gently remove the supernatant (flow-through or non-bound) and save for analysis if desired. 17. Wash the beads with 3 × 500 μl 25 mM Ammonium bicarbonate, pH 8, by gently vortexing the beads to resuspend during each wash step (see Note 9). 18. Wash the beads with 2 × 500 μl of LC-MS grade water, by gently vortexing the beads to resuspend during each wash step (see Note 10).

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19. Elute bound antigen by adding 100 μl elution buffer (0.5 % formic acid/30 % acetonitrile), and incubating for 5 min at room temperature with periodic gentle vortexing (see Note 11). 20. Collect the beads on a magnetic stand and transfer the elution to a fresh low-binding microcentrifuge tube. 21. Dry the elution in a speed vac. 22. Proceed to mass spec sample prep (Subheading 3.3) and/or resuspend in SDS-PAGE sample buffer for Western blot analysis/ validation. 3.3 Mass Spec Sample Prep and Data Analysis

The proteins present in the IP elution fractions are typically denatured, reduced, alkylated, and digested into peptides prior to detection by mass spectrometry, because peptides fractionate, ionize, and fragment more efficiently than intact proteins. In addition, the resulting peptide mass spectra are easier to interpret during the protein identification process. Because the protein amount in the IP eluent is small, an in-solution digestion workflow is preferred as it minimizes sample loss, is fast and is amenable to high-throughput and automation. Samples are first denatured in a strong chaotrope such as urea to facilitate complete digestion. Disulfide bonds are reduced with tris(2-carboxyethyl) phosphine (TCEP) or dithiothreitol (DTT) and bond reformation is prevented by alkylating free sulfhydryl groups with iodoacetamide. The fully denatured proteins are then digested with trypsin, although other proteases and protease combinations can be used (e.g. chymotrypsin, Glu-C, Lys-C). The resulting peptides are applied to C-18 tips or columns to remove salts and buffers and to concentrate the sample prior to analysis by nano or capillary reversed-phase liquid chromatography (LC), electrospray ionization, and tandem mass spectrometry (MS/MS). The peptides are identified by sequence database searching using the search algorithm SEQUEST or Mascot. The significance of each peptide and protein identification is estimated using the software tools, Thermo Scientific™ Proteome Discoverer™ 1.4 and Scaffold 4.0 software to assess percent sequence coverage, spectral counts, and post-translational modification (PTMs). 1. Suspend dried samples in 10 μl denaturation buffer (see Note 17). 2. Add 10 μl of 10 mM TCEP to each sample and incubate at 37 °C for 30 min. Centrifuge briefly to collect condensate to bottom of tube. 3. To each 20 μl sample of reduced protein add 0.83 μl of 0.5 M iodoacetamide solution (final concentration 20 mM). Incubate at room temperature for 30 min in the dark (cover sample tubes with foil OR place samples in a drawer). 4. Add 45 μl of 50 mM Tris–HCl, pH 8.0 to dilute urea concentration to Advanced Crop > Define Crop Area, see Fig. 2). 4. Create a new experiment (File > New Experiment). Select both of the cropped images. Move through the next set of screens by clicking “Next.” This will select the default options. 5. Follow the instructions on the Spot Detection Parameter Wizard. Increase the sensitivity to maximize the number of spots identified (see Fig. 3). 6. Select Proceed. In the next dialog box, select the cropped image of the stained gel as the Master gel. Select “Add unmatched spots from all gel images to the Master gel.” This will create an artificial image containing all of the detected spots from both the stained blot and the immunodetected Western blot (see Fig. 4). 7. Choose default values by clicking Next through the successive dialog boxes. The software will calculate matches and display the Master image, the blot images and an Experiment Summary with match rates (see Fig. 5). “Match Rate 2” for the Western blot image is the ratio of the number of protein spots recognized by the antibody to the total number of proteins detectable in the sample (see Fig. 6). 8. Choose View > Spot > Show Crosshairs or press F5 to show the detected spots (see Fig. 7). 9. Show matched spots by selecting elect Analyze > Analysis Set Manager…, then Create and Matching (see Fig. 8). The resulting images will have identified spots marked with a colored “+” and matched spots marked with an “x” in a different color. This will aid in the determination of whether or not a spot identification or match is valid (see Fig. 9).

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Fig. 2 “Advanced Crop” of the 2-D images

2-D Western Blotting for Anti-HCP Evaluation

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Fig. 3 Spot detection

10. Determine a more accurate match rate by deleting incorrectly identified spots. These can often be visually identified near the edges of the image. In this case shown in Fig. 10, a series of spots along the top edge of the blot image probably do not correspond to real proteins. Delete spurious spots by selecting Edit > Spot > Remove Spot. The spots are erased by clicking on the spot or drawing a box around a set of spots. This operation should be performed both on the Raw 2-D image and on the Master. 11. Select Edit > Experiment Summary to display the final corrected match rate (see Fig. 11). 12. If desired, the match may be further refined by manually adding and deleting spots. Spot matches may be systematically verified using the Spot Review Tool (Analyze > Spot Review see Fig. 12).

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Fig. 4 Choose Master Gel

Fig. 5 Spot matching

Fig. 6 Match rate

Fig. 7 Detected spots

Fig. 8 Show matched spots

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Fig. 9 Matched spots

Fig. 10 Erase misidentified spots


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Fig. 11 Corrected match rate

Fig. 12 Spot review

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Notes 1. Solutions containing urea are prepared immediately before use. 2. It is convenient to use a concentrated stock solution of Bromophenol Blue, but this dye will not dissolve in unbuffered water. It is therefore prepared in Tris base solution. 3. Other pH ranges may be used. pH 3–10 NL (non-linear) was chosen for this experiment because it covers most of the range of protein pI’s. This non-linear gradient is flatter in the region of the gradient where most proteins are found, thereby spreading them out and allowing resolution of more individual proteins. 4. Equilibration buffer is highly viscous and it may be difficult to stir efficiently. Allow several hours for the urea to dissolve into the liquid components. 5. The amount of Equilibration solution with DTT or iodoacetamide may be scaled up or down depending on the number of IPG strips run. These solutions may be prepared from aliquoted Equilibration buffer (see Subheading 2.3 step 3). Frozen tubes of Equilibration buffer should be allowed to thaw slowly in a beaker of water (do not heat).

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6. This solution contains SDS and is highly prone to boiling over. It should be microwaved at low power with constant monitoring. When the solution starts to boil over, turn off the power, remove from the microwave oven and swirl before returning to the microwave oven. Repeat until the agarose is fully dissolved. Aliquots of agarose sealing solution will solidify and can be re-melted before use in a 95 °C heat block or by immersion in boiling water. 7. This stock solution gives 1× TBS with a composition of 20 mM Tris-Cl pH 7.5, 500 mM NaCl. 8. Non-fat dry milk generally gives high-stringency, low background immunodetection. Alternative blocking reagents may be substituted. 9. This primary detection antibody is from goat, necessitating the use of an anti-goat secondary detection reagent. 10. This imaging instrument allows both fluorescent and chemiluminescent imaging at the same resolution with the same camera. This simplifies alignment and matching of the total protein (fluorescent) image and immunodetected (chemiluminescent) image. 11. This method could be applied to cell culture supernatant and cellular lysate prepared from other cell types. 12. Alternative cell culture conditions may be employed, but cells should be grown in protein-free, defined medium so that no proteins that do not derive from CHO cells will be found in the cell culture medium. Suspension culturing will result in a higher concentration of secreted protein in the growth medium and will require less concentration to produce a sample to be analyzed by 2-D electrophoresis. 13. Concentrated cell culture supernatant contains interfering substances that will prevent effective analysis by 2-D electrophoresis. The 2-D Cleanup Kit is a convenient means for removing these substances and further concentrating the sample by selective precipitation. Other methods, such as acetone precipitation [8], have been used successfully. The dry protein pellet may be slow to dissolve. Brief sonication will accelerate the process but do not allow the material to heat up. 14. Protein is most accurately quantitated in solution lacking detergent or reductant, so concentrated protein samples are first prepared without these additives. The detergent (CHAPS) and reductant (DTT) are supplied by the solution in which the sample is diluted prior to first-dimension SDS. 15. IEF may be conducted in either 8 M urea or in 7 M urea, 2 M thiourea. The urea/thiourea mixture results in better protein solubilization, particularly of hydrophobic proteins, but urea


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alone generally results in a cleaner, higher resolution 2-D pattern [9]. In this experiment, the secreted proteins, which are generally very soluble, are separated in a solution containing urea alone. Cell lysate protein, which contains more hydrophobic proteins, is separated in a solution containing a urea/ thiourea mixture. 16. Each of the sample solution components has a specific role in promoting high resolution IEF separation. Urea and thiourea are protein denaturants that promote the complete unfolding of polypeptide chains so that all ionizable groups are exposed to the solution. CHAPS is a detergent that prevents protein aggregation and promotes solubility. DTT is a thiol reductant that breaks disulfide bonds within and between polypeptide chains, and maintains proteins in a fully reduced state. Carrier ampholytes enhance protein solubility by minimizing protein aggregation due to charge–charge interactions. Bromophenol blue is a tracking dye. Its inclusion is not necessary, but clearance of the dye from the IPG strip during electrophoresis provides a visual confirmation that electrical current is being delivered to the IPG strip appropriately. 17. Each sample is run in replicate and separated on two seconddimension gels, one of which is stained and the other blotted. 18. Recombinant therapeutics can be expressed either as secreted proteins recovered from the culture medium or as cellular proteins that are recovered from cell lysates. The HCPs found in either case are expected to differ from each other. Cell culture medium, however, is expected to contain some proteins that are released from cells that lyse during growth and transfer. This experiment analyzes reactivity of an antibody directed against secreted HCP in both a secreted protein- and cell lysate-derived samples (see Figs. 13 and 14). 19. Handle IPG strips from the ends using forceps. 20. The sample is applied to the entire length of the immobilized pH gradient by rehydration into a dry IPG strip. As voltage is applied, each peptide in the sample focuses to a narrow zone in the pH gradient corresponding to its pI. 21. Mineral oil prevents evaporation of the sample solution. 22. Additional details of the sample application and IEF procedure may be found in the user manual for the PROTEAN i12 IEF System. 23. It will take at least 10 min for the agarose sealing solution to melt. It is best to start this prior to equilibrating the IPG strips for the second dimension. 24. Although size standards are not strictly required for this application, they may be applied to the gel at this stage if desired.

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Fig. 13 CHO cell lysate probed with anti-CHO HCP: Stained gels, stained blot, and Western blot. CHO cell lysate was separated in duplicate by 2-D electrophoresis. One gel was stained with Oriole Fluorescent gel stain (a). The second gel was transferred to PVDF membrane and stained with Oriole following transfer (b). The PVDF membrane was stained with SYPRO Ruby Blot Stain to visualize total protein (c). Following staining, the blot was immunodetected with anti-CHO HCP and visualized by chemiluminescence (d)

Colored standards (e.g. Precision Plus Protein WesternC Standard, Bio-Rad) can serve to verify electrophoresis and transfer and to orient the blotting membrane and determine which side contains transferred protein. Apply 5 μl of standards solution into the standards well using a syringe or gel loading pipet tip. 25. The 10 min low voltage step is to allow gradual transfer of the proteins from the IPG strip to the second-dimension gel. This reduces vertical streaking and gives a higher resolution second-dimension separation. 26. The fluorescent stain used is resistant to photobleaching and shielding the staining reaction from normal room light should not be necessary. If the gel is under bright light, or in sunlight, the staining tray may be covered with aluminum foil. 27. The gel is delicate and easily torn. Handle the gel wearing gloves and avoid touching the gel anywhere besides the edges and corners. The gel is best held by the bottom corners. Place the gel on the anode stack and membrane by draping the gel over the membrane starting from the top of the gel. Once the


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Fig. 14 CHO cell secreted protein probed with anti-CHO HCP: Stained gels, stained blot and Western blot. CHO cell secreted protein was separated in duplicate by 2-D electrophoresis. One gel was stained with Oriole Fluorescent gel stain (a). The second gel was transferred to PVDF membrane and stained with Oriole following transfer (b). The PVDF membrane was stained with SYPRO Ruby Blot Stain to visualize total protein (c). Following staining, the blot was immunodetected with anti-CHO HCP and visualized by chemiluminescence (d)

gel is in place on the membrane, it should not be lifted again or moved. 28. The gel is stained following transfer to verify completeness of transfer. 29. At this stage, the membrane may be allowed to dry and can be stored for several weeks at room temperature between sheets of blotting paper. 30. SYPRO Ruby Protein Blot Stain is a sensitive fluorescent stain for proteins blotted onto PVDF or nitrocellulose membranes. It is used in this application because it does not interfere with subsequent immunodetection [10]. 31. Alternatively, this step may be carried out for 1 h at room temperature. 32. The substrate solution should form a puddle that covers the blot. Ensure that the surface of the blot is completely covered with substrate solution. 33. The default settings for capture of chemiluminescence images take advantage of binning in order to shorten exposure times

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at the expense of image resolution. In order to capture images at sufficient resolution for analysis of 2-D gels, and to generate images at the same resolution as the fluorescent images generated for total protein, binning is turned off, or set to 1 × 1. This may require exposure times of several minutes. References 1. Eaton LC (1995) Host cell contaminant protein assay development for recombinant biopharmaceuticals. J Chromatogr A 705: 105–114 2. Dagoussat N, Haeuw J-F, Robillard V, Damien F, Libon C, Corvaïa N, Lawny F, Nguyen TN, Bonnefoy J-Y, Beck A (2001) Development of a quantitative assay for residual host cell proteins in a recombinant subunit vaccine against human respiratory syncitial virus. J Immunol Methods 251:151–159 3. Wan M, Wang Y, Rabideau S, Moreadith R, Schrimsher J, Conn G (2002) An enzymelinked immunosorbent assay for host cell protein contaminants in recombinant PEGylated staphylokinase mutant SY161. J Pharm Biomed Anal 28:953–963 4. Savino E, Hu B, Sellers J, Sobjak A, Majewski N, Fenton S, Yang T-Y (2011) Development of an in-house, process-specific ELISA for detecting HCP in a therapeutic antibody, Part 1. Bioprocess Int 9:38–45 5. Wang X, Hunter AK, Mozier NM (2009) Host cell proteins in biologics development: identification, quantitation and risk assessment. Biotechnol Bioeng 103:446–458

6. Zhu-Shimoni J, Yu C, Nishihara J, Wong RM, Gunawan F, Lin M, Krawitz DC, Liu P, Wandoval W, Vanderlaan M (2014) Host cell protein testing by ELISA and the use of orthogonal methods. Biotechnol Bioeng 111: 2367–2379 7. Tscheliessnig AL, Konrath J, Bates R, Jungbauer A (2013) Host cell protein analysis in therapeutic protein bioprocessing—methods and applications. Biotechnol J 8:655–670 8. Valente KN, Schaefer AK, Kempton HR, Lenhoff AM, Lee KH (2014) Recovery of Chinese hamster ovary host cell proteins for proteomic analysis. Biotechnol J 9:87–99 9. Rabilloud T, Adessi C, Giraudel A, Lunardi J (1997) Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18:307–316 10. Berggren K, Steinberg TH, Lauber WM, Carroll JA, Lopez MF, Cernokalskaya E, Zieske L, Diwu Z, Haugland RP, Patton WF (1999) A luminescent ruthenium complex for ultrasensitive detection of proteins immobilized on membrane supports. Anal Biochem 276: 129–143

Chapter 29 Free Flow Electrophoresis for Separation of Native Membrane Protein Complexes Lutz Andreas Eichacker, Gerhard Weber, Ute Sukop-Köppel, and Robert Wildgruber Abstract This chapter describes the technology of free flow electrophoresis (FFE) and protocols to separate membrane protein complexes for proteome analysis. FFE is a highly versatile technology applied in the field of protein analysis. It is superior to native PAGE due to its fast continuous processing of sample at high resolution. Additionally, the dynamic separation range from ions, peptides, to proteins, protein complexes, up to organelles, and whole cells makes it the method of choice in the analysis of proteins. FFE is carried out in an aqueous medium without inducing any solid matrix, such as acrylamide, so that it simplifies the analysis of protein complexes for the downstream analysis. Here, we describe the novel zone electrophoresis interval method (IZE-FFE) for separation of protein complexes from the thylakoid membrane of Arabidopsis thaliana by charge only. Protein complexes isolated by IZE FFE were characterized according to molecular weight by Blue Native PAGE and were proteins stained with coomassie. Key words Free flow electrophoresis, Thylakoid membrane, Protein complexes, Blue Native PAGE

Abbreviations BN FFE HPMC iZE PAGE ZE

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Blue Native PAGE Free flow electrophoresis Hydroxy-propyl-methylcellulose Interval zone electrophoresis Polyacrylamide gel electrophoresis Zone electrophoresis

Introduction The molecular mass ranges of thylakoid membrane protein complexes that regulate biogenetic processes cover several orders of magnitude. Isolation and characterization of protein complexes and analysis by single particle electrotomography or structure


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determination by crystallization requires the biological material in a liquid environment after separation. In the last years, native PAGE has become an established laboratory standard for the separation of protein complexes [1–4]. The system provides excellent resolution, but it provides the isolated protein complexes in a gel environment and high-resolution separation of complexes takes 16 hours. The problem of releasing the protein complexes from the gel is solvable; however, the additional steps compromise complex stability and require time before the molecules can be analyzed further. In general, there are three problems with native gels that can be overcome by using free-flow electrophoresis. Disassembly of protein complexes at the liquid–gel interface is avoided, an upper limit for entering of complexes into the gel is circumvented, and separation of the complexes is tremendously speeded up. Why is it essential to overcome the gel-based limitations? The liquid/gel junction confronts proteins with a deleterious molecular barrier for entry into the molecular sieve network and the integrity of many complexes that fulfill the entry size requirement is compromised. For supercomplex analysis, the molecular mass entry limit of highresolution polyacrylamide gels according to molecular mass standards appears at about 1.5 MDa; although separation of assemblies with a molecular mass of about 10 MDa has been claimed [5, 6]. Size exclusion limits an analysis of functional cell structures that operate in supramolecular arrangements. Analysis of cellular complexity and characterization of the biochemical dynamics requires to analyze complex arrangements over large molecular mass differences top down, e.g. from the cell to the organelle, to the membrane, and the polysomal/cotranslational level in one experimental system. A barrier-free analysis strategy is therefore essential to analyze the dynamics of regulatory cell processes. Finally, many complexes are labile and disintegrate during prolonged separation. It is therefore essential to keep the time between sample preparation and analysis of the cell components as short as possible. Free-flow electrophoresis offers a solution to overcome the constraints of native PAGE. As the name implies, the technique is gel-free. Using interval zone electrophoresis, separation of membrane protein complexes of interest has been achieved within about 6 min (Fig. 1). Separation is based on the native charge of the complexes and resolution is achieved by mobility differences in defined pH zones. The separation technology can be automated and has been used for analytical and preparative tasks [6–8]. Separated protein fractions are pooled in microtiter plates and hence are directly accessible in the liquid state [8]. Steps for analysis of the free-flow fractions, including native and SDS-PAGE, spectroscopy, mass spectrometry, and single particle analysis have successfully been conducted and optimized recently.

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Fig. 1 FFE-separated protein complexes visualized after native BN-PAGE. Thylakoid membranes corresponding to 300 μg Chl were solubilized in 300 μL detergent mixture 1 and 2. Supernatant 2 was applied to Free Flow Electrophoresis. The FFE fractions containing the solubilized protein complexes were then concentrated and subsequently loaded onto native BN-PAGE. Chl-protein complexes and standard proteins (kDa) were stained with colloidal Coomassie G250 and detected by white light scanning

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Materials

2.1 Preparation of Thylakoid Membranes (Rosette Leaves from Arabidopsis thaliana)

All buffers should be freshly prepared and stored at 4 °C. 1. Extraction buffer: 25 mM Tricine-NaOH, pH 7.8, 330 mM sorbitol, 1 mM Na-EDTA, 10 mM KCl, 0.15 % (w/v) BSA, 4 mM sodium ascorbate. 2. Lysis buffer: 10 mM Tricine-NaOH, pH 7.8, 5 mM MgCl2, 10 mM NaF. 3. Washing Buffer: 25 mM Tricine-NaOH, pH 7.8, 100 mM sorbitol, 5 mM MgCl2·, 10 mM KCl, 10 mM NaF. 4. Storage buffer (TMKGS): 10 % (v/v) glycerol, 25 mM TricineNaOH, pH 7.8, 100 mM sorbitol, 5 mM MgCl2·, 10 mM KCl.

2.2 Solubilization Buffers for the Membranes

1. DDM solution: 195.84 mM (10 % w/v) n-dodecyl-β-Dmaltoside. Store at −20 °C. 2. DIG solution: 81.34 mM (10 % w/v) digitonin (see Note 1). Store at −20 °C. 3. Solubilization solution 1: 16 mM Digitonin, 250 mM Sucrose, 50 % v/v TMKGS. 4. Solubilization solution 2: 8 mM Digitonin, 8 mM n-dodecylβ-D-maltoside, 250 mM Sucrose, 50 % v/v TMKGS.

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FFE Media

1. Anodic stabilization medium (inlet 1): 100 mM HCl, 50 mM Formic acid, 50 mM Hydroxy-isobutyric acid (HIBA), 250 mM Sucrose, adjust with BisTris to pH 3.8–4.1. 2. Separation medium 1 (inlets 2, 3, 4): 10 mM HIBA, 250 mM Sucrose, 0.1 % Digitonin, adjust with BisTris to pH 5.4. 3. Separation medium 2 (inlets 5, 6): 10 mM HIBA, 250 mM Sucrose, 0.1 % Digitonin, adjust with BisTris to pH 6.2. 4. Separation medium 3 (inlet 7): 10 mM HIBA, 250 mM Sucrose, 5 mM NaCl, 0.1 % Digitonin, adjust with BisTris to pH 7.0. 5. Separation medium 4 (inlet 8): 10 mM HIBA, 250 mM Sucrose, 0.1 % Digitonin, adjust with BisTris to pH 7.0. 6. Cathodic stabilization medium (inlet 9): 150 mM HIBA, 375 mM Imidazol, 250 mM Sucrose. 7. Counterflow medium: 250 mM Sucrose, 50 mM BisTris, 20 mM N-(1,1-Dimethyl-2-hydroxyethyl)-3-amino-2hydroxypropanesulfonic acid (AMPSO), resulting in pH 8.0. 8. Anodic electrode medium: 100 mM sulfuric acid. 9. Cathodic electrode medium: 100 mM sodium hydroxide, 200 mM Glycine, resulting in pH 10. 10. FFE System: FFE advanced, FFE Service GmbH, Munich, Germany. 11. Viva-Spin 100,000 MWCO tubes at 10 °C (Viva-Products, Inc., Littleton, MA).

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BlueNative-PAGE

1. Thylakoid membranes from Arabidopsis thaliana: 1 μg/μL total Chlorophyll, frozen in liquid N2. Stored at −80 °C. 2. Cathode buffer: 50 mM Tricine, 50 mM BisTris, resulting in pH 6.8, 0.02 % Coomassie G250. Assemble as 10× concentrate. Store at 4 °C (see Notes 2 and 3). 3. Anode buffer: 50 mM Tricine, 50 mM Bis-Tris, resulting in pH 6.8. Assemble as 10× concentrate. Store at 4 °C (see Note 3). 4. Electrophoresis gel: 3–12 % Bis-Tris-Gels, 100 × 100 mm. 5. Electrophoresis chamber: vertical slab gel electrophoresis unit (e.g. mini cell from Novex). 6. Power supply.

2.5 Analysis of Thylakoid Membrane Complexes

1. Molecular weight standard: NativeMark (Invitrogen). 2. White light scanning: Epson Perfection V750, transmission mode scanner. 3. Colloidal Coomassie stain: 0.12 % (w/v) Coomassie G250, 20 % (v/v) Methanol, 10 % (v/v) Phosphoric acid, 10 % Ammonium sulfate [9].


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Methods The FFE system performs electrophoretic separations in patented and well-defined separation solutions without the use of a solid (e.g. polyacrylamide) matrix. The gel-free basis enables the highresolution separation of charged or chargeable entities like cell organelles, protein complexes and proteins, or peptides on a fast, preparative, and continuous basis. Fluid-phase separation is operated in the three modes zone electrophoresis (ZE), including the novel iZE method, isoelectric focusing (IEF), and isotachophoresis. The system enriches low-abundant proteins with high reproducibility. The sample is applied using a peristaltic pump and is induced into a separation chamber consisting of two parallel plates. Under laminar flow, the sample is transported within a thin (0.2–0.4 mm) film of aqueous medium formed between the two plates. The plates are bordered by two electrodes that generate a high-voltage electric field perpendicular to the laminar flow. Charged particles like ions, peptides, proteins, organelles, membrane fragments, or whole cells that are deflected in this electric field are separated and seamlessly fractionated into microtiter plates. The iZE method described in this chapter delivers excellent resolution for the separation of protein complexes and membrane proteins. Compared to IEF separations, media require no addition of polymers like Hydroxy-ethyl-propyl-cellulose (HPMC) to suppress electroendosmotic flow. This enables an easy concentration of the separated samples by ultra-filtration and ensures highest compatibility for any downstream technology like, e.g. native and SDS-PAGE, HPLC, MS, and ELISA. The protocol describes how protein assemblies from the thylakoid membrane of Arabidopsis thaliana can be prepared for iZEFFE separation. Separation by iZE-FFE is based on the charge of the protein assemblies. The resolution of the iZE-FFE protocol is visualized using blue native PAGE as a native PAGE approach. In general, blue-native PAGE is intended to separate protein complexes in a state that reflects the physiologically relevant functional state of the protein assembly [4]. It is therefore advisable to minimize protein complex degradation through sample preparation on ice and fast electrophoresis at 4 °C. Solubilization of thylakoid membranes resulted in the formation of membrane assemblies of chlorophyll-binding thylakoid membrane protein complexes that were charge separated by iZE-FFE within 6 min. The composition of the FFE fractions was analyzed in a BN-PAGE approach. Here, one to several protein bands have been determined per FFE fraction. Unique protein compositions were identified per every second FFE fraction that was loaded for BN-PAGE separation. Especially, five distinctive maxima were determined after the FFE fractionation (Fig. 1, fractions 35, 37, 49, 53 and 59). BN-PAGE

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lanes and proteins bands were analyzed further by denaturing SDS-PAGE and proteins and complexes were identified by mass spectrometry. Data revealed that iZ-FFE was perfectly suited to separate the highly complex array of membrane protein complexes from the photosynthetic thylakoid membrane. Protein complexes and subcomplexes were well detected upon Coomassie staining and white light scanning (Fig. 1). Detailed characterization of protein complexes and subcomplexes is presently under investigation. Separation of the protein complexes by native PAGE is based on a sieving of the complexes by the acrylamide concentrationdependent pore size of the gel. The method presented here corresponds to a 3–12 % (v/v) linear gradient separating gel (Fig. 1). The gradient gel facilitated a separation of molecular weight standards in the molecular weight range of 66–1,500 kDa (Fig. 1). 3.1 Arabidopsis thaliana Sample Preparation

1. Arabidopsis thaliana plants are grown on soil for 3–4 weeks in a growth chamber with ambient white light of 100 μmol/m2/s at a light/dark cycle of 8 h-light/16 h-dark. 2. Homogenize leaves 4 × 4 s in 150 mL of ice-cold extraction buffer. 3. Filter homogenized leaves through one layer of Miracloth and collect through a funnel in 3 × 50 mL conical tubes. 4. Centrifuge for 3 min at 1,800 × g. Collect the pellet and discard the supernatant. 5. Gently resuspend the pellet in 5 mL extraction buffer using a paintbrush. Wash the brush in 25 mL extraction buffer. Redistribute into two conical tubes and fill to 50 mL with extraction buffer. 6. Centrifuge for 3 min at 1,800 × g. Collect the pellet and discard the supernatant. 7. Gently resuspend each pellet in 5 mL of lysis buffer with a paintbrush. 8. Combine resuspended material in one conical tube and fill to 50 mL with lysis buffer. 9. Divide sample into two Sorvall HB4 tubes and fill to 25 mL with lysis buffer. 10. Incubate sample for 5 min in the dark on ice. 11. Centrifuge for 5 min at 6,000 × g. 12. Supernatant = chloroplast stroma. Purify stroma from residual thylakoids by centrifugation at 12,000 × g for 30 min. 13. Pellet = chloroplast thylakoid membrane. 14. Gently resuspend thylakoid membrane pellets from both tubes in 5 mL of wash buffer with a paintbrush and collect in one tube.


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15. Collect thylakoids by centrifugation for 5 min at 5,900 × g. 16. Discard the supernatant and keep the pelleted thylakoids. Resuspend thylakoids in 3 mL wash buffer using paintbrush. 17. Transfer into glass homogenizer and lever piston three times. 18. Transfer into 5 mL plastic tube. 19. Clean homogenizer with 2 mL of wash buffer and homogenize 2× and collect in 5 mL tube. 20. Measure the concentration of chlorophyll. Transfer 10 μL thylakoid membrane extract into 990 μL 80 % Acetone at 4 °C. Determine absorbance at 652 nm. Calculate the concentration of Chl (μg/μL). 21. For storage of thylakoid membranes determine Chl concentration and dilute in storage buffer to a concentration of 1 μg Chl/μL. 22. Freeze aliquots of 100 μL in liquid N2 and store tubes at −80 °C. 3.2 Solubilization of Thylakoid Membranes from Arabidopsis thaliana for Native FFE

1. Use thylakoid membranes corresponding to 300 μg Chl. 2. Concentrate thylakoid membranes by centrifugation at 7,500 × g and 4 °C for 10 min and discard supernatant. 3. Solubilize in 300 μL of detergent mixture 1 for 10 min at 10 °C (see Note 1). 4. Separate non-solubilized material 25,000 × g and 10 °C for 10 min.

by

centrifugation

at

5. Transfer the supernatant 1 containing the solubilized protein complexes to a new micro tube. 6. Solubilize the pellet in 300 μL of detergent mixture 2 for 10 min at 10 °C. 7. Separate non-solubilized material 25,000 × g and 10 °C for 10 min.

by

centrifugation

at

8. Transfer the supernatant 2 containing the solubilized protein complexes to a new micro tube. 9. Load supernatant of the different solubilizations onto FFE (see Fig. 1 for separation of supernatant 2) (see Note 2). 3.3

FFE Method

A native separation buffer and the novel interval Zone Electrophoresis protocol is used for protein complex separation. FFE is conducted at 10 °C using the following conditions: The experiments run in a horizontal position of the separation chamber using a 0.2 mm spacer. The voltage is adjusted to 1,600 V which results in a current of ~85 mA. The residence time in the separation chamber is approximately 4.5 min per interval. Fractions are collected in polypropylene microtiter plates numbered from 1 (anode) through 96 (cathode).

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1. Place the sample to the ice cooled sample holder on the FFE system (see Notes 3–7). 2. Clean interior surface of chamber: move separation chamber to an upright position and open the pairs of chamber clamps simultaneously. Use the following sequence for cleaning (H2Odist ≤ 18 nΩ/20 μS):water– isopropanol–petrolether–isopropanol–water (see Note 8). 3. Apply the spacer (0.2 mm) without covering the inlets, apply the electrode membrane (soaked in Glycerol/Isopropanol, mix 1:1) and the filter paper (0.3 mm), soaked in distilled water onto the membrane. 4. Close separation chamber: lift the Plexiglas side of the chamber to meet the mirrored side, first fasten central and adjacent clamps loosely (release chamber clamps by two turns each). Then fasten central and adjacent clamps tight, close top and bottom clamps, place fractionation plate on top of fractionation housing, finally make sure that all inlets of media and counterflow are free and are not covered by the spacer. 5. Fill separation chamber with degassed dist. water: open (lift) valves at the top of the chamber to degas the chamber, fasten wedge clamps (I1–I9) of the media pump and place tubes in reservoir filled with H2Odist, switch on medium pump on control board. When the chamber is totally flushed and free of air add counter flow by closing wedge clamps on pump. 6. Set up the Electrode Buffer, Stabilization Buffers and counter flow buffer and separation buffers. 7. Connect anode (+) and cathode (−) electrolytes, connect plastic cover and start electrode pump. 8. Adjust the media flow rate to 120 mL/h and start the media flow, wait for 15 min to flush the buffers in. 9. Adjust the voltage to 1,600 V, set the current to 150 mA. 10. Set the sample flow rate to 4,000 μL/h. 11. Set the media pump flow rate to 16.4 digits. 12. Turn on media pump. 13. Turn on sample pump and flush the sample in for 70 s, then switch the sample pump to backwards direction for 2 s and turn off the sample pump. 14. Wait 20 additional seconds, and then turn on the high voltage. 15. Wait 30 s, and then adjust the media pump to 3.6 units. 16. Wait 4 min, then turn the high voltage off and elute the separated samples with a media pump speed of 28.8 units.


423

17. After 100 s the sample can be collected with a MTP or deep well plate, therefore place the MTP in the fraction collection device and collect for 120 s. 18. If more separated sample is needed, repeat the steps 8–17. 3.4 Concentration of FFE Fractions

3.5 Electrophoresis of Protein Complexes from FFE Fractions on BN PAGE

For native LN- and BN-PAGE analysis, a volume of 200 μL of each IZE fraction was concentrated by centrifugation using Viva-Spin 100,000 MWCO tubes at 10 °C (Viva-Products, Inc., Littleton, MA) according to the manufacturer’s protocol until a volume of 25 μL was retained. 1. Prepare cathode-, anode- and gel-media for BN-PAGE-gels. Store at 4 °C. 2. Label the position of the wells on the outer cassette with a felt pen and remove the comb carefully from the native gel (see Note 9). 3. Assemble cassette sandwich within electrophoretic apparatus. 4. Fill in cathode buffer into the upper buffer chamber. 5. Rinse the wells with cathode buffer (see Note 10). 6. Underlay the samples into the wells using a micro syringe or disposable pipette tips (see Note 11). 7. Fill anode buffer in the lower buffer chamber until electrode is immersed. 8. Place the gel chamber in a Styrofoam box, use 2 minus 20 °C cooling pads to maintain low temperature during separation. 9. Complete assembly of electrophoresis unit and connect to power supply. 10. Set the power supply to limit voltage. Use 35 V constant. Set mA and W to maximal values. Run overnight for highest resolution. Apply power to electrophoretic set (see Note 12). 11. Stop electrophoresis when the front composed of Ponceau S/chlorophyll has reached the bottom of the separating gel (see Note 13). 12. For further analysis of protein complexes like absorbance or mass spectroscopy measurements open sandwich and cut chlorophyll gel bands or stain proteins before further processing of sample of interest.

4

Notes 1. Digitonin is dissolved by heating the solution to 100 °C. After heating, Digitonin can immediately be mixed with other detergents. The rest of the detergent mixture can be stored at −20 °C. Always reheat solutions before usage.

424


2. Depending on the organism and membrane that should be solubilized, different detergent concentrations and mixtures need to be tested individually. 3. Turbidity of protein samples is an indication of insufficient solubility and/or protein precipitation. Protein samples may not be turbid; otherwise the resolution of the electrophoretic separation will be poor, turbid protein samples have to be cleared by centrifugation prior to the FFE separation. 4. The salt concentration of samples may not exceed 25 mM. If the conductivity of the sample due to the salt concentration is too high, it has to be desalted or diluted to reach 5,000) and reflectron (m/z < 5,000) mode of TOF-MS. 7. The microwave oven has 2,450 MHz frequency and the maximum power is 700 W. A thermocouple probe was used to measure the temperatures of sample solutions prior to and after the microwave experiments. 2.1 Functionalization of Nanomaterials

1. Quantum dots (QDs) were synthesized by the well described procedures in the literature [14]. Similarly, the unmodified gold nanoparticles (Au NPs) were prepared by using NaBH4 as a reducing agent. After preparation of bare QDs and Au NPs, dopamine dithiocarbamate was attached on the surfaces of QDs by the described procedures [14, 15]. Briefly, 0.549 mM dopamine (104.1 mg), 0.549 mM carbon disulfide (40 μL), and triethylamine (5 μL) were taken in a 2.0 mL glass vial and sonicated for 5 min at room temperature. The formed dopamine dithiocarbamate was added into a 25 mL of nanomaterials (QDs and Au NPs) and then stirred for 15 min at room temperature. Finally, the product was washed with ethanol to remove reactants and unbound dopamine dithiocarbamate. 2. Vanillin, ethanol, acetic acid, and 4-aminothiophenol were used for the synthesis of (4-mercaptophenyliminomethyl)-2methoxyphenol (Schiff base 3). Then, the bare Au NPs dispersed aqueous solution was added to the toluene solution containing 1.66 mM of 4-mercaptophenyliminomethyl-2methoxyphenol and the solution was stirred vigorously for 5 h. The Au NP was successfully transferred to the organic phase (toluene), as indicated by the color change from light yellow to dark brown [16]. 3. 400 μL of MPA was transferred into a 250 mL round bottom flask containing 15 mL of deionized water. To this, 2 mL of 0.01 M cadmium nitrate was added dropwisely under N2 pressure with constant stirring for 1 h. The pH of the solution was adjusted to 9 by adding ammonium hydroxide solution. 2.5 mL of 0.008 M sodium sulfide solution was quickly added to the above solution at 96 °C and solution was stirred for 2 h. The obtained clean green-yellowish CdS QDs solution was stored at 4 °C until further use [17]. Similarly, HgTe nanostructures were prepared according to a previously described procedure [7].

484

Suresh Kumar Kailasa and Hui-Fen Wu

2.2 Preparation of Hydrophobic Nanomaterials

1. AgNO3, Se powders and octadecylamine (ODA), octadecanethiol (ODT) and 11-mercaptoundecanoic acid (MUA) were used for the synthesis of functionalized silver selinide nanoparticles (Ag2Se NPs) [18]. The formed NPs were washed with ethanol and then dispersed in toluene using an ultrasonicator for further use. 2. Surface-modified BaTiO3 NPs were synthesized according to the literature procedure [19]. Briefly, BaTiO3 NPs (0.5 g) surfaces were hydroxylated by refluxing with 200 mL of H2O2 for 4 h at 106 °C. The hydroxylated BaTiO3 NPs were filtered and then washed with deionized water for two times. The hydroxylated BaTiO3 NPs were directly added into 250 mL round bottom flask that contained 50 mL of 50 % (v/v) aqueous alcohol along with 0.5 (w/v) of 12-hydroxy octadecanoic acid (HOA). The reaction mixture was heated at 90 °C under vigorous stirring for 3 h. The reaction mixture was cooled to room temperature and HOA-modified BaTiO3 NPs were treated with 1 N HCl until the pH became 5–6 in order to agglomerate the nanopowder and to facilitate NPs filtration. The surface-modified BaTiO3 NPs were washed with deionized water and acetone in sequence, several times. The surfacemodified BaTiO3 NPs were dried in vacuum oven for 24 h and then dispersed in toluene by ultrasonication for 10 min.

2.3

3

Bacteria Growth

Escherichia coli (12570) standard culture bacteria were obtained from Bioresource Collection and Research Center, Taiwan. Glassware and media were subjected to autoclave at 15 lbs of pressure for 15 min prior to bacteria culture. One colony of E. coli was carefully taken up from a freshly prepared 24 h old streak plate culture using a sterile loop. The collected bacteria were cultured on Luria Broth Agar (LBA) plates for 24 h at 37 °C. All microbiological experiments were performed in a Biosafety Level 1 cabinet [19].

Methods

3.1 Functionalized Nanomaterials as Matrices for Peptides and Proteins

1. The stock solutions of peptides (Leu-enk, Met-enk, and HW6) were diluted further for working concentrations. An aliquot 900 μL of the peptide solutions were taken in a 1 mL polyethylene vial and 0.1 M HCl or NaOH was added to control pH of the solution. 100 μL of (0.5–6.0 μM) Mn2+-doped ZnS semiconductor nanoparticles solution was added and vortexed for 30 min with speed of 86 × g and then centrifuged the vials at 6,797 × g. The sample solutions (1 μL) were pipetted onto a stainless steel target plate and allowed to air-dry for 10 min before the MALDI-MS analysis ([20], Note 1).

Proteomic Profiling by Nanomaterials

485

2. 750 μL of standard solution of analytes (proteins or peptides) were taken into a 1 mL polyethylene vial. 250 μL of known concentration of CdS QDs solution was added and pH of the test solution was controlled by adding 0.1 M HCl or NaOH. The sample vials were vortexed for 30 min at 86 × g and then 1 μL of the above test solution was placed on the target plate for MALDI-TOF-MS analysis [17]. 3. The prepared HgTe nanostructures were used as concentration and desorption/ionization matrixes. Sample solutions (1 mL) containing a protein, HgTe nanostructures (0.08×), and 300 mM ammonium citrate, pH 5.0 were equilibrated at ambient temperature for 30 min. The mixtures were then centrifuged (3,823 × g, 10 min), and all the supernatants were removed. The pellets were resuspended in ammonium citrate buffer (20 μL), and the sample solutions (1 μL) were pipetted onto a stainless steel target plate and allowed to air-dry for 30 min prior to SALDI-MS measurement ([7], Note 2). 4. Furthermore, HgTe nanostructures are used as the matrix for the investigation of two protein − protein complexes: α1-antitrypsin − trypsin and IgG − protein G. Trypsin (1.7 − 15 μM), α1-antitrypsin (5 μM), and Zn2+ ions (0.1–4 μM) were equilibrated in ammonium citrate solutions (20–200 mM, pH 8.0, 50 μL) at 25 °C for 1 h; protein G (2–20 μM), IgG (10 μM), and Zn2+ ions (0.1–4 μM) were equilibrated in ammonium citrate solutions (20 mM, pH 5.0, 50 μL) at 25 °C for 1 h. Equal volumes of the HgTe nanostructures (4×, 10 μL) in the presence of 0.1 or 1 % Brij 76 and one of the protein mixtures (10 μL) were mixed by vortexing for 1 min. Aliquots (1.0 μL) of the mixtures were pipetted onto a stainless-steel 96-well MALDI target and dried in air at room temperature for 30 min prior to SALDI-MS analysis [21]. 3.2 MicrowaveAssisted Tryptic Digestion

1. Eppendorf tubes are used to perform microwave-assisted tryptic digestions of BSA, cytochrome c, α-casein, and non-fat milk. First, 50 μM of the above proteins were dissolved in 50 mM of NH4HCO3 buffer (pH 8.3) containing 8.0 M of urea. To this, 2.5 μL of 1.8 M CaCl2 was added and then reduced by the addition of 5.0 μL of 50 mM DTT for few minutes. The final concentration of each protein (18.0 μM; 250 μL) was prepared by using 50 mM of NH4HCO3 and urea was maintained below 2 M. To these solutions, trypsin was added at an enzyme-toprotein ratio at 1:40 (w/w) for the microwave-assisted digestion. The above eppendorf tubes were placed in plastic rack and then microwave irradiation was performed by applying microwave power at 210 W for 50 s. After completion of microwave irradiation, sample solutions were subjected to cool for few minutes prior to enrichment procedures with nanomaterials as

486


affinity probes. All the microwave tryptic-digested samples were diluted to a certain concentration with 50 % (v/v) ACN that contained 0.1 % of TFA. 2. 500 μL of non-fat milk (50-folds of dilution) was denatured by adding 50 mM of NH4HCO3 and 8 M of urea and the sample vials were vortexed for a few minutes at room temperature. To this, 100 μL of 50 mM DTT and 5.0 μL of 1.8 M CaCl2 were added and then 200 μL of trypsin (mg/mL) was added. The microwave irradiation was performed at 210 W for 50 s. The sample vials were taken out and cooled for a few minutes and digested proteins were enriched by the aforesaid procedures. 3.3 Nanomaterials as Affinity Probes for the Enrichment of Digested Proteins

A solution of the suspended nanomaterials (BaTiO3 NPs—50 μL, 12 mg/mL; QDs-100 μL, 100 nM) was added to 150 μL of a microwave tryptic digest sample. The mixture was vortexed for 30 min at room temperature. Then, nanomaterials-conjugated digested proteins were isolated by subjecting to centrifugation at 2,655 × g for 10 min at room temperature. The obtained supernatant solution was removed using micropipette (see Note 3). The nanomaterials-conjugated digested proteins (phosphoproteins and non-phosphoproteins) were collected at the bottom of sample vial. To avoid washing procedures, directly 1 μL of 0.2 M 2,5-DHB solution was mixed with 1.0 μL of nanomaterials-conjugated digested protein solution and then 0.5 μL of nanomaterialsconjugated digested protein solution was placed on the MALDI target for MALDI-MS analysis ([14], Note 4). The developed workflow was successfully applied for the identification of phosphopeptides in milk ([21], Note 5).

3.4 Nanomaterials as Extracting and Concentrating Probes for Enrichment of Peptides and Proteins

The nanomaterials-based SDME experiments were performed by the following procedures. The target analytes with desired concentrations were spiked into a glass vial filled with 1 mL sample solution. A 10 μL of microsyringe was used to draw 1 μL of toluene containing the modified nanomaterials. The microsyringe was inserted into the sample solution through a PTFE-coated silicon septum of screw cap of a glass vial. As soon as the sample was extracted into the 1 μL microdroplet of organic solvent (the modified nanomaterials prepared in toluene), the microdroplet was drawn back into the microsyringe and then directly placed the 1 μL microdroplet onto the target plates for subsequent MALDI-MS analysis ([16], Note 6). Nanomaterials-based LLME: To this, 900 μL of standard peptide mixture (valinomycin—0.40 μM; gramicidin D—2.0 μM) was taken into a 1.0 mL polyethylene vial and then pH of the sample was adjusted by adding 1.0 M of HCl or NaOH. Then, 100 μL of


487

dispersed functionalized Ag2Se NPs (2.0 mg/mL) was added and then vortexed at 86 × g for 30 min. The sample vials were allowed to stand for 3 min to separate dispersed Ag2Se NPs in toluene (organic) and aqueous layers. After that 2 μL of dispersed functionalized Ag2Se NPs conjugated hydrophobic peptides (valinomycin and gramicidin D) was taken by using micropipette and mixed with equal volume of 0.2 M 2,5-DHB. The above sample was allowed to air dry prior to MALDI MS analysis [18]. For hydrophobic proteins in E. coli: 200 μL of cultured E. coli (1.5 × 108 cfu/mL) was taken into a 1.0 mL vial and then 200 μL of HOA-modified BaTiO3 NPs dispersed in toluene was added. The vials were vortexed for 30 min at room temperature. The sample vials were kept on stands for 3 min to separate NPs contained in organic layer and in aqueous layers. Then, 1.0 μL of HOAmodified BaTiO3 NPs-conjugated hydrophobic proteins (upper layer) was mixed with 1.0 μL of 0.5 M SA. Finally, 1.0 μL of above solution was placed on the MALDI target plate, dried at room temperature and then analyzed by MALDI-MS ([19], Note 7). 3.5 Peptides and Proteins Identification by Nanomaterials-Based MALDI-MS

Among different possible bioanalytical approaches, nanomaterialsbased MALDI mass spectrometry-based proteomics is increasingly used to acquire the data important for understanding these processes. For example, Mn2+-doped ZnS-cysteine NPs-based MALDI-MS was successfully used for the analysis of peptide mixture of Leu-enk (3.0 μM), Met-enk (3.0 μM), and HW6 (2.1 μM) (Fig. 1). It can be observed that Mn2+-doped ZnS-cysteine NPsbased MALDI-MS provided remarkable mass spectra for peptide mixture, corresponding to the mass peaks of peptides at m/z 578.2, 595.3, 615.6, 906.6, and 929.5 for [Leu-enk + Na]+, [Metenk + Na]+, [Leu-enk + K]+, [HW6 + H]+, and [HW6 + Na]+, respectively. All ion signals were generated with sodium and potassium adducts while only HW6 peptide was generated as protonated ions. This reason is due to the alkali metals have greater affinity toward the small molecules in MALDI-MS and also due to the nanomaterials as the matrix lacking of the proton sources. Moreover, large proteins were successfully detected in E. coli by using HgTe nanostructures as matrices in MALDI-MS (Fig. 2). The peaks at m/z 10,770, 21,539, 10,728, and 21,454 that correspond to the adducts [zSSAT1 + 2H]2+, [zSSAT1 + H]2+, [zSSAT1 R101A + 2H]2+, and [zSSAT1R101A + H]+, respectively, resulting from the recombinant proteins of zSSAT1 or zSSAT1 R101A under individual plasmids transformed into E. coli strain BL21. Although zSSAT1 and zSSAT1 R101A share 99.5 % amino acid sequence homology and only one amino acid variation, our approach allowed differentiation of them, showing its potential for the rapid detection of proteins. Interestingly, HgTe nanostructuresbased MALDI-MS was effectively detected protein G in both the

488


Fig. 1 MALDI-mass spectra of peptide mixture (Leu-enk, Met-enk, and HW6) using (a) CHCA and (b) Mn2+doped ZnS-cysteine nanoparticles as the matrices. Peak identities: m/z 578.2, 595.3, 615.6, 906.6 and 929.5 which are attributed to the [Leu-enk + Na]+, [Met-enk + Na]+, [Met-enk + K]+, [HW6 + H]+ and [HW6 + Na]+. Reproduced from ref. 20 with permission from Royal Chemical Society

absence and presence of Brij 76, but the protein G − IgG complex was detectable only in the presence of 0.1 % Brij 76 [22]. The signals at m/z 26,023 and 13,019 represent the [protein G + H]+ and [protein G + 2H]2+ adducts. The mass peaks at m/z 74,885, 49,984, 26,023, 13,019, 86,585, and 58,297 correspond to the [IgG + 2H]2+, [IgG + 3H]3+, [protein G + H]+, [protein G + 2H]2+, [IgG+ protein G + 2H]2+, and [IgG + protein G + 3H]3+ adducts (Fig. 3). Using this approach, singly and multiply charged (1+, 2+, 3+, and 6+) adducts of IgG were observed and this approach successfully allowed to detect protein G (m/z 26,023) and IgG (m/z 149,931) at concentrations as low as 2 μM (1 pmol) and 5 μM (2.5 pmol).


489

Fig. 2 Mass spectra of solutions containing HgTe nanostructures and the recombination proteins (0.01 μg/μL) (a) zSSAT1 and (b) zSSAT1 R101A. SALDI-MS was performed in the linear mode. The peaks at m/z 10,770, 21,539, 10,728, and 21,454 represent the adducts [zSSAT1 + 2H]2+, [zSSAT1 + H]+, [zSSAT1R101A + 2H]2+, and [zSSAT1 R101A + H]+, respectively. Reproduced from ref. 7 with permission from The American Chemical Society

The identification and characterization of post-translational modifications (PTMs) of proteins can be easily studied by top-down fragmentation of intact protein ions. However, traditional proteolysis requires 6–12 h and tedious separation procedures are needed for their identification. These are limitations for rapid identification of digested proteins with high selectivity and sensitivity. To overcome this problem, functionalized QDs are used as affinity probes for the enrichment of microwave-tryptic digested BSA in MALDI-MS (Fig. 4). Using this approach, the protein digestion time was drastically reduced from 12 h to 50 s with good sequence coverage. From the obtained MALDI mass spectrum, 11 proteolytic peptides (B1, B3, B6, B8, B9, B16, B21, B22, B24, B25, and B26) were identified from microwave tryptic digest of BSA with low sequence coverage of 26 %. In order to obtain more efficient microwave tryptic digestion of proteins, QDs-DDTC were used as concentrating probes for the enrichment of microwave-tryptic digest of BSA without any pretreatment procedure. As a result, many digest fragments of BSA (35 peptides from B1 to B35) were observed with high signal-to-noise ratios by using QDs-DDTC as affinity probes.

Fig. 3 Mass spectra of IgG, protein G, and their complexes, recorded through SALDI-MS using HgTe nanostructures (a) in the absence and (b and c) in the presence of 0.1 % Brij 76. The samples were prepared in 20 mM ammonium citrate (pH 5.0) containing 1 μM Zn(II). The signals at m/z 149,931, 74,885, 49,984, 24,997, 26,023, 13,019, 86,585, and 58,297 represent the adducts [IgG + H]+, [IgG + 2H]2+, [IgG + 3H]3+, [IgG + 6H]6+, [protein G + H]+, [protein G + 2H]2+, [IgG + protein G + 2H]2+, and [IgG + protein G + 3H]3+, respectively. Reproduced from ref. 22 with permission from The American Chemical Society

Fig. 4 MALDI-TOF mass spectra of microwave tryptic-digested peptides originated from BSA (6.0 μM) using (a) 2,5-DHB as the conventional matrix and (b) QDs-DDTC as the affinity probes. Reproduced from ref. 14 with permission from Elsevier

491


a 5000

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Fig. 5 (a) Direct MALDI mass spectrum of microwave tryptic digest of α-casein (4.0 × 10−6 M). (b) MALDI mass spectrum of microwave tryptic digest of α-casein using BaTiO3 NPs (12 mg/mL) as enrichment probes. Reproduced from ref. 21 with permission from Springer

These results indicate that the identified proteolytic peptides were matched with the amino acid sequence coverage of 56 % for BSA. Furthermore, 24 phosphopeptides (from α1 to α24) are effectively identified with improved signal intensities using BaTiO3 NPs as enrichment probes from the microwave tryptic digest of α-casein (Fig. 5b). Figure 5 provides remarkable evidence that some of the low abundance phosphopeptides (α12, α16, α18, α19, and α20) are effectively identified using BaTiO3 NPs. The BaTiO3 NPs were used as enrichment probes to concentrate phosphopeptides from non-fat milk. Notably, 21 phosphopeptides (from Mα1 to Mα21) were effectively detected in MALDI-MS using BaTiO3 NPs as the concentrating probes (Table 1). The phosphopeptide peaks of β-casein in non-fat milk are appeared at m/z 1,980.6, 2,350.5 and 2,475.1, which are due to dephosphorylated fragment ions (loss of phosphorylated groups) from the ions at m/z 2,061.2, 2,431.3 and 2,556.2, respectively (data were not shown). The identified phosphopeptides in non-fat milk are listed in Table 1. Functionalized Au NPs were successfully integrated in SDME technique for the extraction of milk proteins in milk. The mass peaks at m/z 9,168.01, 11,498.10, 14,218.10, and 18,394.56 are

492


Table 1 List of identified phosphopeptides by MALDI-MS from the microwave tryptic digest of non-fat milk (50-fold dilution) using BaTiO3 NPs as affinity probes Sr. No

Peak number

Observed m/z

Phosphopeptide sequences

1

Mα1

924.4

DIGpSESTEDQAMEDIK (α-S1/58–73)

2

Mα2

976.2

YKVPQLEIVPNpSAEER (α-S1/119–134)

3

Mα3

1,003.3

NANEEEYSIGpSpSpSEEpSAEVATEEVK (α-S2/61–85)

4

Mα4

1,027.8

DIGpSEpSTEDQAMEDIKQ (α-S1/43–59)

5

Mα5

1,103.5

GNAEGpSpSDEEGKLVIDEPAK (α-S1/180–188)

6

Mα6

1,251.6

TKVIPYVRYL (α-S2/213–222)

7

Mα7

1,266.8

YLGYLEQLLR (α-S1/91–100)

8

Mα8

1,293.7

QMEAEpSIpSpSpSEEIVPNpSVEQ (α-S1/74–93)

9

Mα9

1,337.5

VNELpSKDIGpSEpSTEDQAMEDIK (α-S1/52–73)

10

Mα10

1,383.7

FFVAPFPEVFGK (α-S1/38–49)

11

Mα11

1,593.6

TVDMEpSTEVFTKK (α-S2/153–165)

12

Mα12

1,637.2

YLGYLEQLLRLKK (α-S1/106–118)

13

Mα13

1,641.3

FFVAPFPEVFGKEK (α-S1/38–51)

14

Mα14

1,759.8

HQGLPQEVLNENLLR (α-S1/23–37)

15

Mα15

1,769.3

LYQGPIVLNPWDQVK (α-S2/114–128)

16

Mα16

1,847.4

DIGpSETEDQAMEDIK (α-S1/58–73)

17

Mα17

1,927.6

DIGpSEpSTEDQAMEDIK (α-S1/119–134)

18

Mα18

2,105.0

TDAPSFSDIPNPIGSENSEK (α-S1/189–208)

19

Mα19

2,235.2

HPIKHQGLPQEVLNENLLR (α-S1-(19–37))

20

Mα20

2,618.7

NTMEHVpSpSpSEESIIpSQETYK (α-S1/17–36)

21

Mα21

2,678.0

VNELpSKDIGpSEpSTEDQAMEDIK (α-S1/52–73)

Reproduced from ref. 21 with permission from Springer. pS refers to phosphorylated serine unit

corresponded to proteoso pep. PP81 [1], γ3-casein [2], α-lactoalbumin [3], and β-lactoglobulin [4], respectively [16]. To investigate the potential applications of functionalized nanomaterials in LLME for extraction and preconcentration of target proteins, HOA-modified BaTiO3 NPs used as extracting and preconcentrating probes for LLME of hydrophobic proteins in E. coli prior to their identification by MALDI-MS. Figure 6 displays the MALDI mass spectrum of identified hydrophobic proteins in E. coli by using HOA-modified BaTiO3 NPs-assisted LLME coupled with MALDI-MS. Using this approach, 14 hydrophobic proteins were successfully extracted and preconcentrated by using HOA-modified


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Fig. 6 MALDI mass spectra of identified hydrophobic proteins in E. coli by using (a) HOA-modified BaTiO3 NPsassisted LLME along with SA (0.5 M) as the matrix and (b) SA (0.5 M) as the matrix without HOA-modified BaTiO3 NPs. Reproduced from ref. 19 with permission from Elsevier

BaTiO3 NPs as hydrophobic affinity probes. Importantly, the signal intensities of hydrophobic proteins (at m/z 5,090, 5,368, 7,009, 7,173, 7,861, and 8,405) were greatly enhanced (2–13 times) by using HOA-modified BaTiO3 NPs as extracting and preconcentrating probes. The mass peaks at m/z 3,418, 7,185, 10,855, 5,878, and 7,020 Da corresponded to the membrane proteins ecnB (P56549), lpp (P69776), and osmE (P23933) and to hypothetical membrane proteins yifL (P39166) and ygdI (P65292), respectively. Similarly, the mass peak at m/z 8,888 corresponded to acetyl-acyl carrier protein (ydhI; acetyl-ACP, P0A6A8; acetylation of the phosphopantetheine sulfur). Evidently, five lipoproteins (ecnB, lpp, osmE, yifL, ygdI) and water-insoluble ATPase proteolipid (at m/z 8,282; atpL, P68699) were effectively extracted, preconcentrated and then identified by using HOA-modified BaTiO3 NPs-LLME coupled with MALDI-MS. Unfortunately, only one peak (at m/z 8,405) is generated with good signal intensity, the remaining peaks [2–7, 11, 12] were not well resolved and some of the peaks [1, 8, 10, 23] were not generated by using SA as the matrix. These results indicated that HOA-modified BaTiO3 NPs have showed high affinity toward hydrophobic proteins of E. coli, since HOA molecules on the surfaces of BaTiO3 NPs have played a fundamental key role for the efficient LLME of hydrophobic proteins in E. coli [19].

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Notes 1. Nanomaterials exhibited high surface area, good photostability, which results to improve the solubility in aqueous media and their conjugation with biomolecules. 2. The mass limit when SALDI-MS was used with HgTe nanostructures as matrixes reaches up to 150,000 Da (IgG), much higher than those obtained using other kinds of nanomaterials. The HgTe nanostructures-based MALDI-MS allowed to detect weak drug–protein complexes (BSA/Y and hCAI-ACZ complexes). Importantly, HgTe nanostructures allow the analyses of proteins and their complexes under mild conditions and with greater tolerance toward salts. 3. BaTiO3 NPs are good candidatures for effective enrichment of phosphopeptides from microwave tryptic digest of α-casein. Moreover, after centrifuging of BaTiO3 NPs–phosphopeptides conjugates from microwave tryptic digests, the trapped phosphopeptides are directly identified by MALDI-MS without further washing procedure step. The entire procedure was completed within 60 min. 4. The high degree of protein digestion was observed by using following microwave conditions (microwave power: 210 W and microwave irradiation time: 50 s) and trypsin-to-protein ratio (1:30, w/w) was used for the digestion of cytochrome c, lysozyme, and BSA proteins in the present study. 5. The BaTiO3 NPs acted as concentrating probes for phosphopeptides from microwave tryptic digest of α-casein, since trace levels of phosphopeptides (α12, α16, α18, α19, and α20) are effectively enriched and identified without non-phosphorylated peaks. It confirms that surfaces of BaTiO2 NPs have high capability to adsorb phosphopeptides from microwave tryptic digest of α-casein. It can be proved that BaTiO3 NPs have welldispersed in solution which can improve signal intensities, S/N ratio and allowed unambiguous trapping of low abundance phosphopeptides (Fig. 2). 6. The binary matrix approach applying Au NP-SDME microdroplets which can be homogeneously mixed with the organic matrix (CHCA or SA) to form homogeneous crystals and thus favorable for sensitive detection of peptides and proteins at low concentration in MALDI-MS. In addition, the Au NP-SDME can also serve as multifunctional nanoprobes for ionization, enrichment, preconcentration, and desalting purposes for a variety of peptides and proteins. Moreover, the sample can be directly deposited onto the MALDI target plates and directly sent for MALDI-MS analysis without the need for any further washing steps or elution processes.


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7. HOA-functionalized BaTiO3 NPs have attained hydrophobic nature by the functionalization of BaTiO3 NPs surfaces with HOA (17-alkyl chains), which facilitates to enhance the analytes extraction and preconcentration through hydrophobic interactions. The use of HOA in MALDI-MS highly improves its performance by acting as affinity probes and possibly as a “buffer” in laser energy transfer to enhance the desorption/ ionization of hydrophobic proteins.

Acknowledgement The authors greatly acknowledge Elsevier, Royal Chemical Society, Springer, and American Chemical Society for giving copyright permission to reuse figures and some of the text for this chapter. References 1. Godovac-Zimmermann J, Brown LR (2001) Perspectives for mass spectrometry and functional proteomics. Mass Spectrom Rev 20:1–57 2. Chiang C-K, Chen W-T, Chang H-T (2011) Nanoparticle-based mass spectrometry for the analysis of biomolecules. Chem Soc Rev 40: 1269–1281 3. Zhu Z-J, Rotello VM, Vachet RW (2009) Engineered nanoparticle surfaces for improved mass spectrometric analyses. Analyst 134: 2183–2188 4. Kailasa SK, Cheng K-H, Wu H-F (2013) Semiconductor nanomaterials-based fluorescence spectroscopic and matrix-assisted laser desorption/ionization (MALDI) mass spectrometric approaches to proteome analysis. Materials 6:5763–5795 5. Kailasa SK, Kiran K, Wu H-F (2008) Comparison of ZnS semiconductor nanoparticles capped with various functional groups as the matrix and affinity probes for rapid analysis of cyclodextrins and proteins in surface-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal Chem 80:9681–9688 6. Kailasa SK, Wu H-F (2011) Semiconductor cadmium sulphide nanoparticles as matrices for peptides and as co-matrices for the analysis of large proteins in matrix-assisted laser desorption/ionization reflectron and linear time-offlight mass spectrometry. Rapid Commun Mass Spectrom 25:271–280 7. Chiang C-K, Yang Z, Lin Y-W, Chen W-T, Lin H-J, Chang H-T (2010) Detection of proteins and protein-ligand complexes using HgTe

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19. Kailasa SK, Wu H-F (2013) Surface modified BaTiO3 nanoparticles as the matrix for phospholipids and as extracting probes for LLME of hydrophobic proteins in Escherichia coli by MALDI–MS. Talanta 114:283–290 20. Kailasa SK, Wu H-F (2010) Interference free detection for small molecules: probing the Mn2+-doped effect and cysteine capped effect on the ZnS nanoparticles for coccidiostats and peptide analysis in SALDI-TOF MS. Analyst 135:1115–1123 21. Kailasa SK, Wu H-F (2012) Rapid enrichment of phosphopeptides by BaTiO3 nanoparticles after microwave-assisted tryptic digest of phosphoproteins, and their identification by MALDI-MS. Microchim Acta 179:83–90 22. Chen W-T, Chiang C-K, Lee C-H, Chang H-T (2012) Using surface-assist laser desorption/ ionization mass spectrometry to detect proteins and protein–protein complexes. Anal Chem 84:1924–1930 23. López-Ferrer D, Cańas B, Vázquez J, Lodeiro C, Rial-Otero R, Moura I, Capelo JL (2006) Sample treatment for protein identification by mass spectrometry-based techniques. TrAC Trend Anal Chem 25:996–1005

INDEX A Acetone precipitation .......................................................410 Affinity chromatography ................................. 160, 211–222, 307–309, 319, 467, 472 Affinity electrophoresis.....................................................327 Alkylation ..........128, 159, 170, 173, 196–197, 206, 236–239, 241–242, 251–253, 257, 261, 262, 267–269, 272, 273, 278, 282, 312, 316, 371, 372, 375, 468, 473 Amyloid-binding proteins ........................................465–475 Antibody........................................... 24, 26, 30, 33, 121, 131, 132, 135–139, 141–143, 147–149, 196, 279, 285, 286, 291, 298, 301, 325, 327, 331, 338, 343–345, 347, 383, 384, 386–390, 394, 398, 402–411, 430, 437, 438, 442–445, 448–450, 452, 457, 460, 462 APV® Gaulin .......................................................................2

B Bead-based array ......................................................441, 442 Bead beater .......................................................................2, 4 Bead impact ......................................................................2–7 Bead mill ..........................................................................5–7 Benzonase.......................... 337, 358, 360, 366, 444, 446, 457 Bicinchoninic acid (BCA) protein assay ........... 310, 313, 444 BioNeb cell disruption........................................................16 Bis-Tris SDS PAGE gels .......................... 328, 329, 344, 345 Blue native electrophoresis ...............................................419 Bradford protein assay .............................. 296, 396, 398, 472 Branson sonifier................................................................296

C Carbamidomethylation ..................................... 288, 363, 474 Carbamylation ............................................ 30, 271, 318, 365 Carrier ampholytes ............................122, 159, 164, 396, 411 Cell culture CHO-K1 cells ............................................................398 Hep 70.4 cells .............................................................451 LIM1863 cells ............................................ 180, 182, 192 primary skeletal muscle cells .........................................57 SW 480 cells ....................................... 168–170, 173, 176 Cell disruption bead impact methods ..................................................2–7 electromotive field ........................................................18 high pressure batch ................................... 2, 3, 10–13, 19 high pressure flow ..................................... 2, 3, 13–16, 19 low pressure ...........................................2, 3, 7, 16, 17, 19

mortar and pestle tissue grinders ....................................9 rotor–stator homogenizer ...........................................7–9 ultrasonic processors ...............................................16–19 Cell lysis LC-MS/MS analysis .................................... 67, 259–273 phosphoprotein analysis..............................................313 Cell wall ....................................................... 10, 73, 228, 263 Chaotrope.................................................................145, 156 Chloroplasts Arabidopsis ......................................... 212–214, 217, 221 pea ...................................................... 212, 214, 216, 221 Chromatography affinity chromatography ............................ 160, 211–222, 276, 306–309, 319, 467, 472 hydrophilic interaction liquid chromatography (HILIC) ................................276, 279–281, 286, 288 hydrophobic interaction chromatography .......... 213, 219, 315, 369 ion exchange chromatography ............ 176, 213, 215, 219 reverse-phase liquid chromatography ................ 145, 154, 168, 174, 176, 369 size exclusion chromatography ......30, 212, 213, 218, 219 CID. See Collision induced dissociation Co-immunoprecipitation..........................................443, 445 Collision induced dissociation (CID) ...................... 288, 300, 302, 357, 359, 363–366 Coomassie Brilliant Blue G-250 ......................................333 Coomassie Brilliant Blue R-250.......................................251 Coomassie staining ...........................................................420 Crosslinking .....................................................................138 Culture medium .......................................... 49, 60, 169, 170, 174, 181, 182, 192, 194, 200–203, 262, 278, 331, 332, 334, 393, 410, 411, 443, 445, 446 CyDye ........................121, 127, 156, 311, 315, 429, 431, 439 Cysteine labeling ......................................................157, 159

D Data normalization stain-free technology ..........................................381–390 western blotting ..................................................381–390 Detergents .................................... 28, 30, 105, 110, 119, 120, 124, 130, 133, 136, 146–148, 168, 203, 250, 254, 257, 265, 270, 311, 334, 410, 411, 417, 421, 423, 424, 428, 451 Dimethylformamide (DMF) ........................... 122, 127, 131, 140, 144, 146, 311, 315, 429, 431

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 1295, DOI 10.1007/978-1-4939-2550-6, © Springer Science+Business Media New York 2015

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PROTEOMIC PROFILING 498 Index Disulfide bridges ..............................................................157 Dithioerythritol (DTE) ....................................................164 Dithiothreitol (DTT) .................................67, 105, 110, 120, 122, 123, 128, 130, 145, 157, 159, 164, 191, 195, 197, 214, 216, 226, 236, 237, 246, 251, 252, 261, 262, 266, 268, 269, 271–273, 278, 296, 298, 311, 312, 358, 361, 371, 372, 375, 396, 397, 400, 409–411, 424, 429, 430, 432, 433, 457, 468, 485, 486 DMEM. See Dulbecco’s modified Eagle medium DMF. See Dimethylformamide Dounce homogenizer .......................................................384 Droplet low pressure nebulizer ...........................................17 Droplet’s impingement .......................................................16 DTT. See Dithiothreitol Dulbecco’s modified Eagle medium (DMEM) ........... 45, 48, 201, 260, 278, 280, 331, 443 Dynamic range compression.......................................99–105 Dyno-Mill ............................................................................6

E Ectosome ..................................................................167, 168 Electron microscopy ............ 94, 173, 175, 183, 191, 196, 206 Electrophoresis affinity electrophoresis ................................................327 blue native PAGE .......................................................419 characterization of extracellular vesicles..............167–168 2-D electrophoresis ................................... 158, 394, 396, 397, 399–400, 410, 412, 413 differential in gel electrophoresis (DIGE) ............. 61, 63, 105, 156–158, 430, 431, 433, 434, 436, 438, 439 free flow electrophoresis .....................................415–424 isoelectric focusing ............................... 51, 119, 122, 154, 156, 158, 176, 231, 293–302, 311, 319, 369, 419, 428, 429, 431, 432, 475 SDS-PAGE ................................................ 267, 312, 429 zone electrophoresis .................................... 416, 419, 421 Electrospray ionization (ESI) .......................... 111, 145, 154, 163, 362, 369 Escherichia coli (E. coli) ............. 7, 73, 452, 484, 487, 492, 493 ESI. See Electrospray ionization Exosome density-gradient separation ................................179–206 electron microscopy ............................ 183, 191, 196, 206 immunoaffinity capture methods........................179–206 ultracentrifugation ..............................................179–206

F Filter aided sample preparation (FASP) .............. 66–70, 237, 240–241, 473 mass spectrometry ..............................................237, 240 Formalin fixed paraffin embedded (FFPE) ...... 109–114, 120 Free flow electrophoresis (FFE) ...............................415–424 French press ..............................................................2, 10–12

G Glycoprotein.............................................................205, 357 enrichment .................................................................357 Glycosidases .....................................................................356 Glycosylation ....................................275–291, 324, 355–357, 366, 367, 456 Gradient centrifugation exosomes .............................................................180, 181 lymphocytes ............................................................33–41 mitochondria .................................................... 83, 93, 94

H Heat stabilization of proteins .......................................21–31 HeLa cells ..........147, 280, 289, 334, 338, 342, 371, 376, 377 HILIC. See Hydrophilic interaction liquid chromatography Homogenizer................................................. 7–9, 13, 15, 78, 80, 82, 89, 91–96, 131, 321, 384, 459, 472 Horseradish peroxidase (HRP)................................. 398, 402 western blotting ..................................................398, 402 Host cell protein antibody coverage ....................................... 398, 403–408 CHO cells ...................................394, 396, 410, 412, 413 2-D electrophoresis ..... 394, 396, 399–400, 410, 412, 413 PDQuest ............................................................398, 403 ProteoMiner ...............................................................100 western blotting ..................................................393–414 Human plasma ...................................................................56 Human serum................................................... 100, 101, 482 Hydrophilic interaction liquid chromatography (HILIC) ................................276, 279–281, 286, 288

I ICPL. See Isotope-coded protein label IEF. See Isoelectric focusing Immobilized metal affinity chromatography (IMAC) ........................................276, 279–281, 284, 289, 291, 306, 308–310, 314 Immobilized pH gradient (IPG) ..................... 100, 122, 127, 128, 154, 158, 231, 294, 296, 298, 300–302, 311, 315, 319, 370, 371, 373, 375, 377, 396, 399–401, 411, 412, 429, 430, 432, 436 Immunoaffinity ...........................33, 137, 139, 168, 179–207 Immunoassay ...................................57, 60, 62, 100, 441–452 multiplexing ........................................................441–452 Immunoblotting .......................................118, 124, 204, 206, 227–228, 327, 331, 340, 343 Immunoprecipitation biotin ...........................................137, 141, 143, 147, 148 crosslinking .................................................................138 magnetic beads ....................135, 137, 138, 140, 142–149 mass spectrometry ..............................................135–149 protein AG ................................. 137–140, 143, 147–149

PROTEOMIC PROFILING: METHODS AND PROTOCOLS 499 Index protein–protein interaction .........................................135 streptavidin ................................. 137, 138, 141–142, 147 In-gel digest .............................................118, 119, 129, 236, 238–240, 250, 259–261, 263, 266–270, 325, 331 In-solution digest ............................................ 136, 138, 145, 147, 172–174, 237, 241–242, 260, 266–270 IPG. See Immobilized pH gradient Isoelectric focusing (IEF) .................................. 51, 119, 122, 154, 156, 158, 176, 231, 293–302, 311, 319, 324, 369, 419, 428–432, 475 Isoelectric point (pl) ..........................159, 294, 306, 369, 377 Isotope-coded protein label (ICPL) .........................162–164 ITRAQ..................................................... 272, 290, 357, 429

L Label-free proteomics....................................... 112–114, 207 Liquid chromatography couples to mass spectrometry (LC-MS) .................... 28, 71, 72, 140, 141, 143, 144, 48, 149, 160–163, 236, 262, 369–378, 469, 474, 475 Loading buffer........................................... 73, 148, 216, 220, 227, 278, 279, 282–286, 290, 291 SDS-PAGE ........................................................227, 229 Lowry protein assay ..........................................................264 Lymphocytes density gradient centrifugation ...............................33–41 magnetic cell sorting ...............................................33–41 sample preparation........................................................34 Lysis buffer .............................................45, 47, 67, 110, 111, 114, 140, 142, 144, 147, 148, 214, 216, 218, 220, 231, 237, 240, 246, 263, 265, 271, 272, 282, 290, 310, 313, 318, 319, 331, 332, 334, 370, 372, 417, 420, 444, 446, 447, 451, 457–459, 468, 473 Lysis solution ....................................................................375 Lysosome ............................................................................76

M Maldi. See Matrix-assisted laser desorption ionization Mass spectrometry (MS) electrospray ionization (ESI) ............................. 111, 112, 145, 154, 163, 240, 299, 301, 362, 369 Maldi .................................................. 163, 246, 479–495 tandem .........................................111, 145, 155, 254, 474 Matrix-assisted laser desorption ionization (Maldi) .................... 22, 163, 236, 245, 272, 479–495 Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS) .............................. 82, 485, 490 Membrane proteins ...................159, 401, 402, 419, 428, 493 complexes ...........................................................415–424 isolation with FFE ..................................... 416, 417 Microarray ........................................................ 455–463, 481 human protein (HuProt) ......................................55–463 Microfluidizer........................................................... 2, 15, 16

Microvesicles ..................... 167, 179, 186, 199, 201, 202, 206 isolation ......................................................................202 Mini Bead Beater .................................................................4 Mitochondria ................................................... 13, 58, 75–96 isolation ..................................................................75–96 Mortar and pestle tissue grinders .........................................9 Mouse brain ......................................277, 294, 295, 297, 468 isolation of synaptosomes ........................... 294, 295, 297 Multiplexing ............................................. 137, 154, 155, 159 Muscle human ..............................................................55–63

N Nanomaterials, MALDI MS....................................479–495 N-terminal sequence analysis ...................................249, 254

O Organelle isolation ........................75, 76, 211, 212, 221, 419 chloroplast, mitochondria ................75, 76, 211, 212, 221

P Paraffin embedding ..........................................................117 Parr cell disruption bomb .......................................10, 12–13 PBS. See Phosphate-buffered saline Peptides desalting.......................................................... 70, 71, 246 fractionation by isoelectric focusing ....................369, 373 glycopeptides .......................277–283, 289, 291, 355–367 phosphopeptides ........................................ 276–283, 288, 289, 291, 294, 300, 302, 324, 326, 486, 491, 492, 494 reversed-phase (RP)-chromatography ............... 154, 160, 168, 176, 249, 369, 474 shotgun proteomics .......................70, 118, 207, 324, 369 Peroxisome .........................................................................76 Phenylmethylsulfonyl fluoride (PMSF) ................... 468, 472 protease inhibitor ................................................468, 472 Phosphatase inhibitor ........................................ 26, 147, 278, 296, 297, 310, 318, 319, 345, 444, 446 Phosphate-buffered saline (PBS) .................... 36, 38–40, 45, 46, 49, 50, 57, 61, 63, 68, 101, 105, 121, 124, 125, 140, 142–144, 147, 169, 170, 174, 188–195, 202, 204, 260, 262, 263, 271, 358, 360, 370, 371, 396, 398, 444–446, 448, 449, 457, 458, 460 Phosphopeptides enrichment .......... 276, 278–280, 282–283, 294, 324, 494 identification............................................... 294, 300, 486 Phosphoproteins affinity electrophoresis ................................................327 enrichment .................................................................326 fluorescent gel staining ...............................................318 Phosphorylation ........................................ 22–27, 31, 56, 58, 114, 147, 275–291, 294, 300–302, 305, 306, 323–351, 442, 443, 447, 448, 456 Phos-tag technology .................................................325–326 Plasma proteins .................................................. 99–101, 104

PROTEOMIC PROFILING 500 Index Polytron ........................................................................2, 8, 9 Posttranslational modifications (PTM) acetylation...........................................................276, 277 glycosylation ....................................... 276, 277, 324, 456 phosphorylation ........... 276, 277, 289, 294, 323, 324, 456 Preadipocytes ..........................................................44–50, 52 Protease inhibitors ................ 50, 84, 110, 114, 214, 216, 260, 270, 278, 290, 310, 318, 358, 360, 444, 457, 468, 472 Protein assays bicinchoninic acid (BCA) assay .................. 310, 313, 444 Bradford assay............................................ 111, 171, 172, 188–189, 193, 203, 264, 296, 298, 396, 398, 472 detergent compatible (DC) assay....................... 120, 124, 130, 370, 372 Lowry assay ................................................................264 NanoDrop .......................................... 237, 238, 241–246 Protein complexes..............168, 415–417, 419–421, 423, 424 Protein depletion ......................................................225–232 Protein digestion alkylation ..................... 170, 261, 262, 268, 269, 272, 273 filter aided sample preparation (FASP) ......... 66–69, 237, 240–241, 473 in-gel digest ........................................118, 119, 129, 236, 238–240, 250, 259–261, 263, 266–270, 325, 331 in-solution digest ....................................... 136, 138, 145, 147, 172–174, 237, 241–242, 260, 266, 268–270 Lys-C .................................................145, 278, 289, 296, 298, 302, 358, 361, 371, 372, 375 OASIS ................. 236, 238–240, 278, 290, 358, 361, 362 reduction.....................................170, 173, 206, 236, 237, 239, 241–242, 261, 262, 266, 268, 272, 278, 282, 375 STAGE-tip................................................. 197, 361, 362 trypsin.................................................111, 112, 114, 123, 137, 141, 145, 146, 163, 168, 170, 173, 176, 197, 236, 237, 239, 240, 246, 253, 261, 262, 267, 268, 278, 282, 290, 358, 361, 371, 372, 375 Protein extraction .................................... 110–112, 117–133, 164, 216, 220, 226, 231, 250, 266, 290, 431 Protein fractionation..........................139, 176, 245, 294, 427 Protein interaction ............................................ 442, 443, 452 Protein labeling dimethyl labeling ........................................ 249–257, 290 fluorescent dyes.................................... 79, 133, 154–159, 309, 315, 318, 324, 428 stable isotopes ............................................. 155, 160–164 (2,4,6-trimethoxyphenyl) phosphonium bromide (TMPP) labeling ..........................................249–257 Protein microarray ............................................ 455, 461, 462 Protein N-termini ....................................................249–257 Protein precipitation acetone ................................................................260, 265 Clean-up kit ................................111, 127, 311, 315, 410 methanol/chloroform......................... 168, 170, 172–174, 176, 260, 265–266, 473 trichloroacetic acid (TCA)...........218, 226, 260, 265, 266

Protein profiling ............................................... 293–302, 481 ProteoMiner dynamic range compression ..................................99–105 high abundance proteins ......................... 60, 99, 100, 201 low abundance proteins ........................................99, 100

R Reducing agents ...............................105, 119, 120, 124, 130, 318, 348, 483 Reversed-phase high performance liquid chromatography (RP-HPLC) ......................................... 160, 198, 278 Ribulose 1,5 Bisphosphate Carboxylase/Oxygenase (RuBisCO) high abundant protein ................................................225 plant leaves .................................................................226 protamine sulfate ................................................225–232 protein depletion ................................................225–232 Rotor/stator homogenizers ...............................................7–9

S Sample cleanup......................................... 118, 157, 164, 307 Sample preparation .....................................22, 24, 44, 57, 61, 65–74, 105, 109, 110, 160, 202, 206, 235–237, 257, 259, 278, 280–281, 294, 307, 311, 315, 331–332, 334, 369–378, 384–385, 396, 398, 416, 419–421, 424, 429–433, 443–447, 467, 473, 481, 482 Secretome analysis ........................................................46, 59 Shotgun proteomics...............65–74, 118, 207, 249, 324, 369 SILAC. See Stable isotope labeling with amino acids in cell culture Silver staining ............................................ 61, 154, 156, 219, 220, 232, 266, 272, 313, 317, 318 Solid phase extraction (SPE) ........................... 238, 243, 244, 358, 361, 371, 372, 374, 376 Sonicator bath sonicator .......................................................68, 398 tip sonicator .................................................. 31, 264, 271 SPE. See Solid phase extraction SPR. See Surface plasmon resonance Stable isotope labeling with amino acids in cell culture (SILAC) ................................135, 160–162, 246, 357 Stain-free technology ...............................................381–390 Surface plasmon resonance (SPR) ................... 326, 466, 467, 469–471, 475 Swiss-Prot ................................................................280, 288 Sypro Ruby ........................ 133, 308, 320, 401, 403, 412, 413

T TCEP. See Tris(2-carboxyethyl) phosphine Tissue grinder ...................................................................2, 9 Tissue homogenization ................................ 1–18, 84, 85, 88 Tris(2-carboxyethyl) phosphine (TCEP)................. 140, 145, 157, 170, 237, 245, 273, 429, 431, 432, 439

PROTEOMIC PROFILING: METHODS AND PROTOCOLS 501 Index Triton X-114 ....................................................................424 Trypsin .............................................26, 57, 67, 70, 111, 112, 114, 123, 126, 137, 141, 145–147, 154, 163, 168, 170, 173, 176, 191, 197, 236, 237, 239–243, 246, 253, 260–262, 267, 268, 270, 271, 278, 282, 288, 290, 296, 298, 299, 301, 302, 340, 357, 358, 360, 361, 371, 372, 375, 468, 473–475, 482, 485, 486, 494 2D fluorescence difference gel electrophoresis (2D-DIGE) ......................................... 118–120, 123, 126–129, 428, 429, 431, 433–438 2D-PAGE ................22, 23, 28, 118, 119, 129, 130, 428, 429

U Ultracentrifugation ........................... 167–172, 175, 179–207 Ultrafiltration ...........................................................181, 182 Urine proteomics ................................................................66

V Vacuum centrifuge ........................................... 123, 126, 192, 197, 240, 245, 252, 253, 267, 270, 283, 285, 286, 291, 359

W Western blotting data normalization ......................................................387 house keeping proteins .......................................381, 382 immunostaining .......................................... 384, 386–388 loading control ....................................................381, 382 stain-free technology ..........................................381–390 Wheaton Potter-Elvehjem tissue grinder .............................9

Z Zone electrophoresis (ZE)........................................ 416, 419