Shotgun Proteomics: Methods and Protocols (Methods in Molecular Biology, 2259) 1071611771, 9781071611777

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Methods in Molecular Biology 2259

Mónica Carrera Jesús Mateos Editors

Shotgun Proteomics Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Shotgun Proteomics Methods and Protocols

Edited by

Mónica Carrera and Jesús Mateos Department of Food Technology, Institute of Marine Research (IIM), Spanish National Research Council (CSIC), Vigo, Spain

Editors Mo´nica Carrera Department of Food Technology Institute of Marine Research (IIM) Spanish National Research Council (CSIC) Vigo, Spain

Jesu´s Mateos Department of Food Technology Institute of Marine Research (IIM) Spanish National Research Council (CSIC) Vigo, Spain

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1177-7 ISBN 978-1-0716-1178-4 (eBook) https://doi.org/10.1007/978-1-0716-1178-4 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface The present edition of Shotgun Proteomics: Methods and Protocols by Springer Nature is a collection of specific shotgun proteomics-based laboratory techniques and applications developed in leading laboratories and proteomics units worldwide. In a shotgun proteomics approach, a mixture of proteins is digested with a protease, and the resulting mixture of peptides is then subjected to single or multiple liquid chromatography separation and analyzed, mainly by mass spectrometry, using different fragmentation approaches. Using bioinformatics and database searching algorithms, putative peptide sequences are then validated and assigned to the identified proteins. This strategy has revolutionized the discipline of proteomics. Currently, high-throughput shotgun proteomics approaches are powerful tools that achieve the identification and quantification of thousands of proteins from a single analysis. The book contains 20 chapters covering a broad range of topics divided in six main parts: (I) shotgun proteomics of extracellular vesicles and subcellular structures (three chapters), (II) shotgun proteomics in non-model organisms (three chapters), (III) clinical proteomics (six chapters), (IV) food proteomics (two chapters), (V) analysis of posttranslational modifications and protein complexes (four chapters), and (VI) data processing and storage (two chapters). Chapters include introductions to their respective topics, lists of the necessary materials and reagents, step-by-step, readily reproducible laboratory protocols, and tips on troubleshooting and avoiding known pitfalls. We are convinced that Shotgun Proteomics: Methods and Protocols will be an ideal and upto-date guide for researchers seeking to understand the proteome of any given biological sample. Vigo, Spain

Monica Carrera Jesu´s Mateos

v

Acknowledgments We would like to express our gratitude to all the coauthors and to Prof. John Walker (Senior Editor) and Monica Suchy (Springer Nature Editor) for their assistance in the preparation of this book.

vii

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

PART I

SHOTGUN PROTEOMICS OF EXTRACELLULAR VESICLES AND SUBCELLULAR STRUCTURES

1 Mesenchymal Stem Cell-Derived Extracellular Vesicle Isolation and Their Protein Cargo Characterization . . . . . . . . . . . . . . . . . . . . . . . . . Miriam Morente-Lopez, Juan A. Fafia´n-Labora, Monica Carrera, Francisco J. de Toro, Concha Gil, Jesu´s Mateos, and Marı´a C. Arufe 2 Clinical Proteomics for the Analysis of Circulating Extracellular Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ´ ngel Garcı´a Maria N. Barrachina and A 3 Utilization of Laser Capture Microdissection Coupled to Mass Spectrometry to Uncover the Proteome of Cellular Protrusions . . . . . . . . . Ana Gordon and Karine Gousset

PART II

3

13

25

SHOTGUN PROTEOMICS OF NON-MODEL ORGANISMS

4 Isolation of Apoplastic Fluid from Woody Plant Leaves: Grapevine and Coffee as a Case Study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ es Andreia Figueiredo and Leonor Guerra-Guimara 5 Shotgun Proteomics for L3 and L4 Anisakis simplex Development Stages. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert Stryin´ski, Jesu´s Mateos, Elz˙bieta Łopien´ska-Biernat, and Monica Carrera 6 A Primer and Guidelines for Shotgun Proteomic Analysis in Non-model Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Angel P. Diz and Paula Sa´nchez-Marı´n

PART III

v vii xiii

49

59

77

CLINICAL PROTEOMICS

7 SWATH-MS Protocols in Human Diseases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Maria del Pilar Chantada-Va´zquez, Marı´a Garcı´a Vence, ˜ ez, and Susana B. Bravo Antonio Serna, Cristina Nu´n 8 Serum Proteomic Profiling in Rheumatoid Arthritis by Antibody Suspension Bead Arrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 Lucı´a Lourido, Rocı´o Paz-Gonza´lez, Cristina Ruiz-Romero, Peter Nilsson, and Francisco J. Blanco

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x

Contents

9 Human Blood Plasma Investigation Employing 2D UPLC-UDMSE Data-Independent Acquisition Proteomics . . . . . . . . . . . . . . . . . . Licia C. Silva-Costa, Bradley J. Smith, Pamela T. Carlson, Gustavo H. M. F. Souza, and Daniel Martins-de-Souza 10 Metaproteomics Analysis of Host–Microbiota Interfaces . . . . . . . . . . . . . . . . . . . . . Sjoerd van der Post and Liisa Arike 11 Mass Spectrometry-Based Analysis of Mycobacterial Single-Colony Proteome. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . John Iradukunda, Tariq Ganief, Jonathan M. Blackburn, and Nelson C. Soares 12 Proteogenomic Approach for Mycobacterium tuberculosis Investigation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julia Bespyatykh, Georgij Arapidi, and Egor Shitikov

PART IV 13

14

153

167

181

191

FOOD PROTEOMICS

Shotgun Proteomics for Food Microorganism Detection . . . . . . . . . . . . . . . . . . . . 205 Ana G. Abril, Ignacio Ortea, Jorge Barros-Vela´zquez, Toma´s G. Villa, and Pilar Calo-Mata Shotgun Proteomics and Protein-Based Bioinformatics for the Characterization of Food-Derived Bioactive Peptides . . . . . . . . . . . . . . . . . . . . 215 Monica Carrera, Manuel Pazos, Santiago P. Aubourg, and Jose´ M. Gallardo

PART V ANALYSIS OF POSTRANSLATIONAL MODIFICATIONS AND PROTEIN COMPLEXES 15

16

17 18

FTSC-Labeling Coupled with 2DE-LC–MS/MS Analysis of Complex Protein Mixtures for Identification and Relative Quantification of Tissue Carbonylome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ oz, and Isabel Medina Lucı´a Me´ndez, Lorena Barros, Silvia Mun Mass Spectrometry-Based Proteomics for Analysis of Hydrophilic Phosphopeptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chia-Feng Tsai, Jeffrey S. Smith, Dylan S. Eiger, Kendall Martin, Tao Liu, Richard D. Smith, Tujin Shi, Sudarshan Rajagopal, and Jon M. Jacobs Rapid Shotgun Phosphoproteomics Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ as, and Daniel Lopez-Ferrer Monica Carrera, Benito Can System-Wide Profiling of Protein Complexes Via Size Exclusion Chromatography–Mass Spectrometry (SEC–MS). . . . . . . . . . . . . . . . . . Andrea Fossati, Fabian Frommelt, Federico Uliana, Claudia Martelli, Matej Vizovisek, Ludovic Gillet, Ben Collins, Matthias Gstaiger, and Ruedi Aebersold

227

247

259

269

Contents

PART VI 19

20

xi

SHOTGUN PROTEOMICS DATA PROCESSING AND STORAGE

Qualitative and Quantitative Shotgun Proteomics Data Analysis from Data-Dependent Acquisition Mass Spectrometry. . . . . . . . . . . . . . . 297 Jesse G. Meyer The jPOST Repository as a Public Data Repository for Shotgun Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309 Yu Watanabe, Akiyasu C. Yoshizawa, Yasushi Ishihama, and Shujiro Okuda

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

323

Contributors ANA G. ABRIL • Departamento de Microbiologı´a y Parasitologı´a, Facultad de Farmacia, Campus Sur 15782, Universidad de Santiago de Compostela, A Corun ˜ a, Spain RUEDI AEBERSOLD • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland; Faculty of Science, University of Zurich, Zurich, Switzerland GEORGIJ ARAPIDI • Federal Research and Clinical Center of Physical-Chemical Medicine, Moscow, Russia LIISA ARIKE • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden MARI´A C. ARUFE • Grupo de Terapia Celular y Medicina Regenerativa. Dpto. de Fisioterapia, Medicina y Ciencias Biome´dicas, Facultad de Ciencias de la Salud, Universidade da Corun˜a, INIBIC-CHUAC, Agrupacion estrate´gica CICA-INIBIC, A Corun˜a, Spain SANTIAGO P. AUBOURG • Department of Food Technology, Spanish National Research Council (CSIC), Institute of Marine Research (IIM), Vigo, Spain MARIA N. BARRACHINA • Platelet Proteomics Group, Center for Research in Molecular Medicine and Chronic Diseases (CIMUS), Universidade Santiago de Compostela, Santiago de Compostela, Spain; Instituto de Investigacion Sanitaria de Santiago (IDIS), Santiago de Compostela, Spain LORENA BARROS • Instituto de Investigaciones Marinas—Consejo Superior de Investigaciones Cientı´ficas (IIM-CSIC), Vigo, Galicia, Spain JORGE BARROS-VELA´ZQUEZ • Departamento de Quı´mica Analı´tica, Nutricion y Bromatologı´a, Area de Tecnologı´a de los Alimentos, Facultad de Veterinaria, Campus Lugo, 27002, Universidad de Santiago de Compostela, A Corun˜a, Spain JULIA BESPYATYKH • Federal Research and Clinical Center of Physical-Chemical Medicine, Moscow, Russia JONATHAN M. BLACKBURN • Division of Chemical & Systems Biology, Department of Integrative Biomedical Sciences, Faculty of Health Sciences, University of Cape Town, Cape Town, South Africa; Institute of Infectious Disease & Molecular Medicine, UCT, Cape Town, South Africa FRANCISCO J. BLANCO • Grupo de Investigacion de Reumatologı´a (GIR), Instituto de Investigacion Biome´dica de A Corun˜a (INIBIC), Complexo Hospitalario Universitario de A Corun˜a (CHUAC), Sergas. Agrupacion CICA-INIBIC, Universidade da Corun˜a (UDC), As Xubias, A Corun ˜ a, Spain; RIER-RED de Inflamacion y Enfermedades Reuma´ticas. Instituto de Salud Carlos III, Madrid, Spain SUSANA B. BRAVO • Proteomic Unit, Instituto de Investigaciones Sanitarias-IDIS, Complejo Hospitalario Universitario de Santiago de Compostela (CHUS), Santiago de Compostela, Spain PILAR CALO-MATA • Departamento de Quı´mica Analı´tica, Nutricion y Bromatologı´a, Area de Tecnologı´a de los Alimentos, Facultad de Veterinaria, Campus Lugo, 27002, Universidad de Santiago de Compostela, A Corun ˜ a, Spain BENITO CAN˜AS • Complutense University of Madrid (UCM), Madrid, Spain

xiii

xiv

Contributors

PAMELA T. CARLSON • Laboratory of Neuroproteomics, Department of Biochemistry and Tissue Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, SP, Brazil MO´NICA CARRERA • Department of Food Technology, Institute of Marine Research (IIM), Spanish National Research Council (CSIC), Vigo, Spain MARIA DEL PILAR CHANTADA-VA´ZQUEZ • Research Unit, Hospital Universitario Lucus Augusti (HULA), Servizo Galego de Sau´de (SERGAS), Lugo, Spain BEN COLLINS • School of Biological Sciences, Queen’s University of Belfast, Belfast, UK FRANCISCO J. DE TORO • Grupo de Terapia Celular y Medicina Regenerativa. Dpto. de Fisioterapia, Medicina y Ciencias Biome´dicas, Facultad de Ciencias de la Salud, Universidade da Corun˜a, INIBIC-CHUAC, Agrupacion estrate´gica CICA-INIBIC, A Corun˜a, Spain ANGEL P. DIZ • Department of Biochemistry, Genetics and Immunology, University of Vigo, Vigo, Spain; Marine Research Center, University of Vigo (CIM-UVIGO), Vigo, Spain DYLAN S. EIGER • Department of Biochemistry, Duke University, Durham, NC, USA; Department of Medicine, Duke University, Durham, NC, USA JUAN A. FAFIA´N-LABORA • Grupo de Terapia Celular y Medicina Regenerativa. Dpto. de Fisioterapia, Medicina y Ciencias Biome´dicas, Facultad de Ciencias de la Salud, Universidade da Corun˜a, INIBIC-CHUAC, Agrupacion estrate´gica CICA-INIBIC, A Corun˜a, Spain ANDREIA FIGUEIREDO • Faculty of Sciences, BioISI – Biosystems & Integrative Sciences Institute, University of Lisboa, Lisbon, Portugal ANDREA FOSSATI • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland FABIAN FROMMELT • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland JOSE´ M. GALLARDO • Department of Food Technology, Spanish National Research Council (CSIC), Institute of Marine Research (IIM), Vigo, Spain TARIQ GANIEF • Division of Chemical & Systems Biology, Department of Integrative Biomedical Sciences, Faculty of Health Sciences, University of Cape Town, Cape Town, South Africa ´ NGEL GARCI´A • Platelet Proteomics Group, Center for Research in Molecular Medicine and A Chronic Diseases (CIMUS), Universidade Santiago de Compostela, Santiago de Compostela, Spain; Instituto de Investigacion Sanitaria de Santiago (IDIS), Santiago de Compostela, Spain MARI´A GARCI´A VENCE • Proteomic Unit, Instituto de Investigaciones Sanitarias-IDIS, Complejo Hospitalario Universitario de Santiago de Compostela (CHUS), Santiago de Compostela, Spain CONCHA GIL • Proteomics Facility-Complutense University and Scientific Park Foundation of Madrid, Madrid, Spain LUDOVIC GILLET • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland ANA GORDON • Biology Department, California State University Fresno, Fresno, CA, USA; Department of Cell Biology, Physiology, and Immunology, University of Cordoba, Cordoba, Spain KARINE GOUSSET • Biology Department, California State University Fresno, Fresno, CA, USA

Contributors

xv

MATTHIAS GSTAIGER • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland LEONOR GUERRA-GUIMARA˜ES • Centro de Investigac¸a˜o das Ferrugens do Cafeeiro, Instituto Superior de Agronomia, Universidade de Lisboa, Oeiras, Portugal; Linking Landscape, Environment, Agriculture and Food, Instituto Superior de Agronomia, Universidade de Lisboa, Lisbon, Portugal JOHN IRADUKUNDA • Division of Chemical & Systems Biology, Department of Integrative Biomedical Sciences, Faculty of Health Sciences, University of Cape Town, Cape Town, South Africa YASUSHI ISHIHAMA • Graduate School of Pharmaceutical Sciences, Kyoto University, Kyoto, Japan JON M. JACOBS • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA TAO LIU • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA DANIEL LOPEZ-FERRER • Thermo Fisher Scientific, San Jose, CA, USA ELZ˙BIETA ŁOPIEN´SKA-BIERNAT • Department of Biochemistry, Faculty of Biology and Biotechnology, University of Warmia and Mazury in Olsztyn, Olsztyn, Poland LUCI´A LOURIDO • Grupo de Investigacion de Reumatologı´a (GIR), Instituto de Investigacion Biome´dica de A Corun˜a (INIBIC), Complexo Hospitalario Universitario de A Corun˜a (CHUAC), Sergas. Agrupacion CICA-INIBIC, Universidade da Corun˜a (UDC), As Xubias, A Corun ˜ a, Spain; RIER-RED de Inflamacion y Enfermedades Reuma´ticas. Instituto de Salud Carlos III, Madrid, Spain CLAUDIA MARTELLI • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland KENDALL MARTIN • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA DANIEL MARTINS-DE-SOUZA • Laboratory of Neuroproteomics, Department of Biochemistry and Tissue Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, SP, Brazil; Experimental Medicine Research Cluster (EMRC), University of Campinas, Campinas, Brazil; Instituto Nacional de Biomarcadores em Neuropsiquiatria, Conselho Nacional de Desenvolvimento Cientı´fico e Tecnologico, Sa˜o Paulo, Brazil; D’Or Institute for Research and Education (IDOR), Sa˜o Paulo, Brazil JESU´S MATEOS • Department of Food Technology, Institute of Marine Research (IIM), Spanish National Research Council (CSIC), Vigo, Spain ISABEL MEDINA • Instituto de Investigaciones Marinas—Consejo Superior de Investigaciones Cientı´ficas (IIM-CSIC), Vigo, Galicia, Spain LUCI´A ME´NDEZ • Instituto de Investigaciones Marinas—Consejo Superior de Investigaciones Cientı´ficas (IIM-CSIC), Vigo, Galicia, Spain JESSE G. MEYER • Department of Biochemistry, Medical College of Wisconsin, Milwaukee, WI, USA MIRIAM MORENTE-LO´PEZ • Grupo de Terapia Celular y Medicina Regenerativa. Dpto. de Fisioterapia, Medicina y Ciencias Biome´dicas, Facultad de Ciencias de la Salud, Universidade da Corun˜a, INIBIC-CHUAC, Agrupacion estrate´gica CICA-INIBIC, A Corun˜a, Spain SILVIA MUN˜OZ • Instituto de Investigaciones Marinas—Consejo Superior de Investigaciones Cientı´ficas (IIM-CSIC), Vigo, Galicia, Spain

xvi

Contributors

PETER NILSSON • Division of Affinity Proteomics, Department of Protein Science, KTH Royal Institute of Technology, SciLifeLab, Stockholm, Sweden CRISTINA NU´N˜EZ • Research Unit, Hospital Universitario Lucus Augusti (HULA), Servizo Galego de Sau´de (SERGAS), Lugo, Spain SHUJIRO OKUDA • Division of Bioinformatics, Niigata University Graduate School of Medical and Dental Sciences, Niigata, Japan IGNACIO ORTEA • Institute Maimonides of Biomedica Investigation of Cordoba, Cordoba, Spain ROCI´O PAZ-GONZA´LEZ • Grupo de Investigacion de Reumatologı´a (GIR), Instituto de Investigacion Biome´dica de A Corun˜a (INIBIC), Complexo Hospitalario Universitario de A Corun˜a (CHUAC), Sergas. Agrupacion CICA-INIBIC, Universidade da Corun˜a (UDC), As Xubias, A Corun ˜ a, Spain MANUEL PAZOS • Department of Food Technology, Spanish National Research Council (CSIC), Institute of Marine Research (IIM), Vigo, Spain SUDARSHAN RAJAGOPAL • Department of Biochemistry, Duke University, Durham, NC, USA; Department of Medicine, Duke University, Durham, NC, USA CRISTINA RUIZ-ROMERO • Grupo de Investigacion de Reumatologı´a (GIR), Instituto de Investigacion Biome´dica de A Corun˜a (INIBIC), Complexo Hospitalario Universitario de A Corun˜a (CHUAC), Sergas. Agrupacion CICA-INIBIC, Universidade da Corun˜a (UDC), As Xubias, A Corun ˜ a, Spain; CIBER Bioingenierı´a, Biomateriales y Nanomedicina (CIBER-BBN). Instituto de Salud Carlos III, Madrid, Spain PAULA SA´NCHEZ-MARI´N • Centro Oceanogra´fico de Vigo, Instituto Espan ˜ ol de Oceanografı´a, Vigo, Spain ANTONIO SERNA • Sciex, Madrid, Spain TUJIN SHI • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA EGOR SHITIKOV • Federal Research and Clinical Center of Physical-Chemical Medicine, Moscow, Russia LICIA C. SILVA-COSTA • Laboratory of Neuroproteomics, Department of Biochemistry and Tissue Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, SP, Brazil BRADLEY J. SMITH • Laboratory of Neuroproteomics, Department of Biochemistry and Tissue Biology, Institute of Biology, University of Campinas (UNICAMP), Campinas, SP, Brazil JEFFREY S. SMITH • Department of Biochemistry, Duke University, Durham, NC, USA; Department of Medicine, Duke University, Durham, NC, USA RICHARD D. SMITH • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA NELSON C. SOARES • College of Pharmacy, Department of Medicinal Chemistry, University of Sharjah, Sharjah, United Arab Emirates; Sharjah Institute for Medical Research, University of Sharjah, Sharjah, United Arab Emirates GUSTAVO H. M. F. SOUZA • Mass Spectrometry Applications & Development, SpectraMass Ltd, Campinas, Sa˜o Paulo, Brazil ROBERT STRYIN´SKI • Department of Biochemistry, Faculty of Biology and Biotechnology, University of Warmia and Mazury in Olsztyn, Olsztyn, Poland CHIA-FENG TSAI • Biological Sciences Division, Pacific Northwest National Laboratory, Richland, WA, USA FEDERICO ULIANA • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland

Contributors

xvii

SJOERD VAN DER POST • Department of Medical Biochemistry and Cell Biology, University of Gothenburg, Gothenburg, Sweden TOMA´S G. VILLA • Departamento de Microbiologı´a y Parasitologı´a, Facultad de Farmacia, Campus Sur 15782, Universidad de Santiago de Compostela, A Corun ˜ a, Spain MATEJ VIZOVISEK • Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, Zu¨rich, Switzerland YU WATANABE • Division of Bioinformatics, Niigata University Graduate School of Medical and Dental Sciences, Niigata, Japan AKIYASU C. YOSHIZAWA • Graduate School of Pharmaceutical Sciences, Kyoto University, Kyoto, Japan

Part I Shotgun Proteomics of Extracellular Vesicles and Subcellular Structures

Chapter 1 Mesenchymal Stem Cell-Derived Extracellular Vesicle Isolation and Their Protein Cargo Characterization Miriam Morente-Lo´pez, Juan A. Fafia´n-Labora, Mo´nica Carrera, Francisco J. de Toro, Concha Gil, Jesu´s Mateos, and Marı´a C. Arufe Abstract In the present protocol, extracellular vesicles (EVs) released from a primary culture of human umbilical cord mesenchymal stem cells (MSCs) were isolated by ultracentrifugation processes, characterized by transmission electron microscopy (TEM) and measured by nanoparticle tracking analysis (NTA). Protein was extracted from EVs using RIPA buffer and then was assessed for integrity. The proteomic content of the total EV protein samples was analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) after labeling by tandem mass tag (TMT). This combined approach allowed the development of an effective strategy to study the protein cargo from MSC-derived EVs. Key words Mesenchymal stem cells, Tandem mass tag, Extracellular vesicles, Transmission electron microscopy

1

Introduction During the last decade, extracellular vesicles (EVs) have gained significant interest due to their role in paracrine responses and intercellular communication. EVs are derived from the endosomal system to form multivesicular structures, which fuse with the plasma membrane and then are released to the extracellular space [1]. EVs reach recipient cells in the local environment (paracrine mode) or are transported to distant tissues via the circulation system (endocrine mode). Numerous investigations have revealed an important role of EVs in intercellular communication under both normal and pathological conditions [2–5]. To study the function of EVs, it is essential to characterize their molecular content, which specifically reflects the phenotype of the cells-of-origin. Comprehensive proteomic analysis has demonstrated that EVs derived from mesenchymal stem cells (MSCs) are packaged with significantly higher fractions of specific protein compared with their cells-of-origin, indicating a regulation of their contents, which

Mo´nica Carrera and Jesu´s Mateos (eds.), Shotgun Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 2259, https://doi.org/10.1007/978-1-0716-1178-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Miriam Morente-Lo´pez et al.

could provide a molecular basis for their distinct functional properties [6]. This protocol describes the steps to process human umbilical cord stromal MSC-derived EVs for subsequent analyses. The focus was the isolation of EVs from culture medium by serial ultracentrifugation and subsequent protein extraction from EVs using RIPA buffer.

2

Materials The cell culture procedures are performed in a class II flow cabinet. All solutions are prepared using ultrapure water (prepared by purifying deionized water to a sensitivity of 18 MΩ cm at 25
C) and analytical-grade reagents. All organic solvent solutions are LC MS-grade. All reagents are prepared and stored at 4
C, and all waste disposal regulations are diligently followed when disposing of waste materials.

2.1 EV Extraction from Culture Medium by Serial Ultracentrifugation

1. Mesenchymal stem cells in culture growing at a 70% confluence. 2. Culture medium Dulbecco’s modified Eagle’s medium (DMEM). 3. Exosome-depleted fetal bovine serum (FBS). 4. Penicillin–streptomycin (P/S). 5. Sodium chloride (NaCl). 6. Filters (0.22 μm). 7. Refrigerated centrifuge 100,000  g.

of

ultracentrifugation

of

soil

8. Culture centrifuge. 9. Ultracentrifuge tubes. 2.2 Particle Number Characterization by Nanoparticle Tracking Analysis (NTA)

1. Syringe of 1-mL Luer slip, Plastipak.

2.3 Morphology Characterization by Transmission Electron Microscopy (TEM)

1. Sodium cacodylate (Na(CH3)2AsO2).

2. PBS (phosphate-buffered saline). 3. Silica microsphere beads. 4. NanoSight LM10.

2. 37% hydrochloric acid (HCl). 3. pH meter GLP 21. 4. Glutaraldehyde (C5H8O2). 5. JEOL JEM-1010 (100 kV) transmission electron microscope.

Extracellular Vesicles from Mesenchymal Stem Cells

2.4 Protein Extraction and Quantification

5

1. RIPA buffer (150 mM sodium chloride (NaCl), 20 mM Tris– HCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS)). 2. Protease inhibitor cocktail. 3. Centrifuge for 1.5-mL tubes. 4. Acetone. 5. Pierce Rapid Gold BCA Protein Assay Kit. 6. Eppendorf® Thermomixer®. 7. Sonicator.

2.5 Protein Digestion and TMT Labeling

1. 1 M triethylammonium bicarbonate, stock solution (TEAB buffer) (C7H17NO3). 2. 200 mM TCEP (Tris(2-carboxyethyl) phosphine hydrochloride (TCEP) (C9H15O6P)) in 0.2 M TEAB. 3. 375 mM iodoacetamide (IAA) (C2H4INO): 1.5 mg IAA + 74 μL of 0.1 M TEAB. 4. Pierce Trypsin Protease MS grade. 5. Culture centrifuge. 6. TMT10plex Mass Tag Labeling Kit. 7. Eppendorf® Thermomixer®. 8. Advanced Vortex Mixer ZX3. 9. Acetonitrile (AcN) (CH3CN). 10. Hydroxylamine (HXAM) (H3NO).

2.6 Peptide BasicReversed-Phase Fractionation

1. High pH Reversed-Phase Peptide Fractionation Kit including 0.1% triethylamine (C6H15N) and spin columns. 2. 0.1% TFA in water. 3. Sonicator. 4. Speed-vac.

2.7 Liquid ChromatographyTandem Mass Spectrometry (LC-MS/ MS)

3

1. Nano-HPLC Bidimensional Dionex UltiMate 3000. 2. The high-resolution Q-Exactive HF.

mass

spectrometer

Thermo

3. The Proteome Discoverer 2.4 search engine.

Methods

3.1 EV Extraction from Culture Medium by Serial Ultracentrifugation

1. Three days before starting the ultracentrifugation process of EV extraction from MSCs in culture, the regular fetal bovine serum is replaced by exosome-depleted fetal bovine serum to guarantee that the EVs do not originate from the fetal bovine serum.

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2. The supernatants are collected and centrifuged at 3000  g for 10 min in a Beckman Coulter Allegra X-22 Centrifuge. The supernatants are then filtered through a 0.22-μm filter and added to new ultracentrifuge tubes. 3. Two serial ultracentrifuges with Hitachi CP100NX are performed. The first one is performed at 100,000  g for 2 h at 4
C. The pellet is then re-dissolved in 15 mL of saline solution. Then, a second centrifugation is done at 100,000  g for 2 h at 4
C, and the last pellet is re-dissolved in 100 μL of lysis buffer or RIPA buffer for protein extraction. 3.2 Particle Number Characterization by Nanoparticle Tracking Analysis (NTA)

The NanoSight LM10 is calibrated using silica microsphere beads. MSC-derived EVs are diluted in PBS in order to obtain a particle number ranging from 108 to 109 particles/μL. At least three repeated measurements of 60 μs are taken per each individual sample, and the mean value is used to determine particle number (Fig. 1a). The movement of each particle in the field of view is measured to generate the average displacement of each particle per unit time, which is calculated using the NTA 3.0.

3.3 Morphology Characterization by Transmission Electron Microscopy (TEM)

1. Re-suspend the EVs in 100 μL of 2.5% glutaraldehyde in 0.1 M cacodylate buffer. This fixation method is the most appropriate for EV observation by TEM to avoid salt precipitation from the medium. 2. Apply the EV suspensions in a metal grid and air-dry. 3. Observe in a JEOL JEM-1010 (100 kV) transmission electron microscope (Fig. 1b).

3.4 Protein Extraction and Quantification

1. Incubate the pellet containing the EVs, which was re-suspended in 100 μL of RIPA buffer, in rotation for 30 min at 4
C. Add 1% protease inhibitor cocktail immediately before use. 2. Centrifuge at 14,000  g for 20 min at 4
C. 3. Add the supernatant to a new tube to which six volumes of acetone was added and precipitate at 20
C for at least 3 h with sporadic gentle shaking (see Note 1). 4. Centrifuge at 14,000  g for 10 min at 4
C. 5. Carefully decant supernatant and briefly air-dry the pellet. Add 100 μL of 0.1 M TEAB. 6. Incubate the pellet in a thermoblock at 25
C with shaking at 1400 rpm for 5 min. 7. Sonicate the pellet for 5 min. Repeat steps 6 and 7 until the pellet is totally dissolved (see Note 2).

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Fig. 1 Characterization of EVs. (a) Representative results from the NTA assay of MSC-derived EV histogram demonstrated that the most of MSC-derived EVs ranged between 50 and 150 nm in diameter in the gauss hood, with median diameters of 133.7 nm. (b) Electron micrographs of MSC-derived EVs from the old and young groups of rats (scale bar ¼ 100 nm)

8. Determine the protein concentration by using a BCA Kit, and verify the protein integrity by SDS-PAGE of 2 μg from each sample and silver staining (see Note 3 and Fig. 2a). 3.5 Protein Digestion and TMT Labeling

1. Precipitate 100 μg of each sample with six volumes of cold (20
C) acetone overnight at 20
C. Spin down the samples at 15,000  g for 15 min at 4
C (see Note 4). 2. Air-dry the pellet for a few seconds and add 100 μL of 0.1 M TEAB.

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Fig. 2 Summary of the proteomic results. (a) Silver staining of a SDS-PAGE. 2 μg of total EV protein from the samples included in the study was loaded per lane. No protein degradation was detected. (b) Number of unique peptides quantified per protein ID. A total of 1848 proteins were identified and quantified, 1141 of them with at least two unique peptides. (c) Percentage of contaminants/human proteins quantified with two or more unique peptides. Only 7% of those proteins were contaminants. (d) Pathway analysis of the most representative biological processes assigned to the proteins identified with two or more unique peptides. Those proteins belong to processes in which EVs are involved, such as exocytosis, secretion, and vesicle-mediated transport

3. Incubate the samples in a thermoblock at 37
C with shaking at 1400 rpm for 5 min and sonicate for 5 min. Repeat until complete dissolution of the protein pellet occurs. Spin. 4. Incubate with 5 μL of 200 mM TCEP in 0.2 TEAB in the thermoblock for 1 h at 55
C. 5. Add 5 μL of 375 mM IAA to each sample and vortex them. Incubate them at room temperature in the dark for 30 min. 6. Add 10 μL of trypsin diluted in 0.1 M TEAB to each sample in a 1:50 (trypsin:total protein) ratio. 7. Vortex and spin and incubate overnight (not more than 16 h) in the thermoblock at 30
C with gentle shaking at 400 rpm. 8. Prepare the TMT10plex Mass Tag Labeling Kit. Let temper at RT for 15 min and briefly spin all the vials. 9. Add 260 μL of AcN to each vial from the commercial TMT10plex Mass Tag Labeling Kit and vortex for 30 s. 10. Incubate for 5 min at room temperature, vortex, and spin.

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11. Add 41 μL of each vial to each corresponding sample and give an average vortex once added to the samples. The leftover reagent could be stored, tightly closed, at 20
C for future experiments (see storage conditions in the manufacturer brochure). 12. Incubate for 1 h at room temperature. 13. Just 5 min before the end of the incubation, prepare 5% HXAM by adding 10 μL of 50% HXAM + 80 μL of H2O (LC-MS) + 10 μL of 1 M TEAB. Vortex, spin, and then add 8 μL of 5% HXAM to the samples. 14. Incubate for 15 min at room temperature. Spin. 15. Transfer the entire contents of each labeling to the same 2-mL vial. The final total volume should be ~1.7 mL containing a total of 1000 μg of labeled peptides if all the channels have been used and 100 μg of total protein per channel (see Note 4) has been processed. 16. Mix gently but vortex and spin. Make aliquots of 100 μg for fractionation and, optionally, take a separate 20 μg aliquot for checking peptide labeling (see Note 5). 17. Dry in speed-vac and proceed to clean-up/fractionation or store at 80
C. 3.6 Peptide BasicReversed-Phase Fractionation

Peptide fractionation is necessary when handling complex samples and allows to improve the number and the sequence coverage of the identified proteins, but at the cost of increasing the number of LC injections. Before fractionation, an optional step is to check by LC-MS/MS the digestion and labeling (see Note 5). 1. Add 300 μL of 0.1% TFA to the dried labeled sample (~100 μg) and vortex/sonicate for 5 min each at 25
C at 1200 rpm. Repeat. 2. In the meantime, prepare the AcN/triethylamine solutions with increasing amounts of AcN (300 μL of each solution per column, see manufacturer brochure). Start with a wash solution of 5% AcN in trimethylamine and increase AcN sequentially by 2.5%, each up to 25%. Make a final elution solution of 50% AcN for very hydrophobic peptides for a total of nine elution steps (see Note 6). 3. Remove the white cap and centrifuge the column for 2 min at 5000  g at 15
C. 4. Discard the liquid. 5. Remove the red cap and add 300 μL of AcN to the column, close the cap and centrifuge for 2 min, 5000  g, 15
C. 6. Discard the liquid and repeat twice. 7. Add 300 μL of 0.1% TFA to the column.

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8. Centrifuge 5000  g for 2 min at 15
C. 9. Discard the liquid and repeat twice. 10. Add 300 μL of sample (~100 μg) in 0.1% TFA. Centrifuge at 3000  g for 2 min at 15
C. 11. Pass the sample back through the column and centrifuge again at 3000  g for 2 min at 15
C. 12. Pick up in the collector and save them again (see Note 7). 13. Add to the column the tubes (H2O-8) already prepared in step 2 from six sample fractionation steps. 14. Centrifuge at 3000  g for 2 min at 15
C. 15. Each eluate (~300 μL) is collected in tubes. Aliquot for peptide quantitation and LC-MS/MS analysis. 16. Dry in speed-vac and proceed to the next step or store at 80
C. 3.7 LC-MS/MS Analysis and Data Processing

Analyze by liquid chromatography-tandem mass spectrometry in a nano Easy-nLC 1000 coupled to a high-resolution mass spectrometer Q-Exactive HF. 1. The peptides were concentrated “online” by reverse-phase chromatography (RP) using an Acclaim PepMap 100 pre-column (20 mm  75 μm ID, C18 of 3 μm particle ˚ pore size). diameter and 100 A 2. Peptides were then separated into a C18 Picofrit reverse-phase analytical column (500 mm  75 μm ID, 2 μm particle diameter, 100 A˚ pore size) with an integrated spray tip, thermostated, operating at a flow of 250 nL/min. 3. The peptides were eluted using a gradient from 2% to 35% of buffer B in 150 min and up to 40% in 10 min. 0.1% FA in water as buffer B and 0.1% formic acid in ACN as buffer A were used. 4. The nano-HPLC is coupled online to the nanoelectric source of the Q-exactive HF mass spectrometer with which the peptides were analyzed. Peptide entry was performed by electrospray ionization using the integrated tip in the analytical column. 5. Data acquisition was carried out with a voltage of 1.8 kV for the electrospray, and the ion transfer tube that guides the ions from the spray to the interior of the mass spectrometer had a temperature of 270
C. 6. The peptides were detected with a resolution of 120,000 in full-scan MS mode in an m/z mass range of 340–1600 Da. 7. The MS/MS data were acquired in the data-dependent acquisition (DDA) mode of the MS. Thus, in each microscan up to 15 precursors with a load of 2+ to 4+ were selected, depending

Extracellular Vesicles from Mesenchymal Stem Cells

11

on their intensity (threshold: 2  104), with dynamic exclusion of 20 s, followed by their isolation with a window width of 2 units of m/z, in a maximum time of 200 ms for fragmentation by HCD (high collision dissociation) with a normalized collision energy (NCE) of 32%. MS/MS spectra were acquired in positive mode. 8. Raw data were processed using Proteome Discoverer 2.4 software. Search was done against the last Uniprot protein databases (human and contaminants, Fig. 2b–d).

4

Notes 1. Acetone precipitation could be prolonged for up to 16 h. 2. If the sample does not completely dissolve, repeat this last step by adding a small amount of TEAB (no more than 20 μL). 3. We strongly recommend checking the protein integrity by SDS-PAGE and silver staining after BCA quantitation. Samples presenting protein degradation should not be included in the TMT study. 4. Scale down the amount of trypsin if necessary. Minimal recommended amount of total protein per channel is 20 μg. In any case, always process the same amount of total protein per channel. 5. LC analysis of peptide fractions is expensive and timeconsuming. Prior to fractionation, we recommend taking a separate 20 μg aliquot and, after C-18 desalting, checking labeling and peptide complexity by LC-MS/MS using a short chromatographic gradient. 6. The number of elution steps can be scaled down depending on the complexity of the sample (observed by SDS-PAGE). Never leave the columns dry and keep wash eluates and sample flowthrough as a precaution in case there is a problem with the binding of the peptides to the C18 resin. 7. We recommend dividing each fraction in three equal parts (~100 μL each). One aliquot could be used for peptide quantitation (using, for instance, the peptide colorimetric quantitation assay); use the other two aliquots for LC-MS/MS analysis of a known amount of peptides to avoid column overloading.

Acknowledgments We thank M. Luisa Herna´ez and M. Dolores Gutie´rrez (from the Proteomics Facility, Complutense University and Scientific Park Foundation of Madrid, Spain) for their excellent MS data acquisition.

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References 1. Barile L, Vassalli G (2017) Exosomes: therapy delivery tools and biomarkers of diseases. Pharmacol Ther 174:63–78. https://doi.org/10. 1016/j.pharmthera.2017.02.020 2. Choi D, Lee TH, Spinelli C, Chennakrishnaiah S, D’Asti E, Rak J (2017) Extracellular vesicle communication pathways as regulatory targets of oncogenic transformation. Semin Cell Dev Biol 67:11–22. https:// doi.org/10.1016/j.semcdb.2017.01.003 3. Chistiakov DA, Orekhov AN, Bobryshev YV (2016) Cardiac extracellular vesicles in normal and infarcted heart. Int J Mol Sci 17(1):63. https://doi.org/10.3390/ijms17010063 4. Han L, Lam EW, Sun Y (2019) Extracellular vesicles in the tumor microenvironment: old

stories, but new tales. Mol Cancer 18(1):59. https://doi.org/10.1186/s12943-019-0980-8 5. D’Anca M, Fenoglio C, Serpente M, Arosio B, Cesari M, Scarpini EA, Galimberti D (2019) Exosome determinants of physiological aging and age-related neurodegenerative diseases. Front Aging Neurosci 11:232. https://doi. org/10.3389/fnagi.2019.00232 6. Yuan O, Lin C, Wagner J, Archard JA, Deng P, Halmai J, Bauer G, Fink KD, Fury B, Perotti NH, Walker JE, Pollock K, Apperson M, Butters J, Belafsky P, Farwell DG, Kuhn M, Nolta J, Anderson JD (2019) Exosomes derived from human primed mesenchymal stem cells induce mitosis and potentiate growth factor secretion. Stem Cells Dev 28(6):398–409. https://doi.org/10.1089/scd.2018.0200

Chapter 2 Clinical Proteomics for the Analysis of Circulating Extracellular Vesicles Maria N. Barrachina and A´ngel Garcı´a Abstract In recent years, technical improvements in proteomics have allowed its rapid application for biomarker discovery, new drug target identification, and the study of disease progression and drug resistance. The clinical potential of circulating extracellular vesicles (EVs) as a source of biomarkers is one of the reasons why several research groups have recently applied proteomics to their study. A large variety of proteomic approaches such as gel-based proteomics and bottom-up and top-down mass spectrometry have been applied to the study of EVs. In this chapter, we will present basic protocols for gel-based and quantitative MS-based approaches applied to the study of EVs. Key words Extracellular vesicles, Platelets, 2D-DIGE, Label-free LC-MS/MS

1

Introduction Over the past decade, the interest in extracellular vesicles (EVs) as a novel source of biomarkers has increased dramatically due to their implication in cell-to-cell communication, cellular activation, inflammation, hemostasis, and disease development and progression [1]. EVs are a heterogeneous population of cell-derived circulating vesicles. However, there is some controversy on nomenclature and sizes of the different types of vesicles. For this reason, the International Society for Extracellular Vesicles (ISEV) has provided some criteria to classify EVs, categorizing them into three groups: microvesicles (MVs), exosomes, and apoptotic bodies [2]. MVs (100–1000 nm) are formed by outward blebbing of the plasma membrane and subsequent fission of plasma membrane blebs. In contrast, exosomes (30–100 nm) are formed by multivesicular bodies being stored on the cytoplasm upon stimulus. The multivesicular bodies fusion with the plasma membrane and release their content (exosomes) to the extracellular environment [3, 4]. Lastly, apoptotic bodies (500–4000 nm) are formed as a result of the induction of cellular apoptosis [5].

Mo´nica Carrera and Jesu´s Mateos (eds.), Shotgun Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 2259, https://doi.org/10.1007/978-1-0716-1178-4_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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So far, extensive research has been carried out to characterize the content of EVs. EVs have a large variety of proteins, sugars, lipids, and genetic material (mRNA, microRNA, rRNA, and tRNA) [6]. The recent development of more robust and sensitive proteomic strategies has allowed a better knowledge of EVs cargo. Indeed, it has been shown that the protein profile of circulating EVs changes when transitioning from health to disease. For this reason, the role of EVs has gained significant interest in the clinical practice as promising targets/sources of biomarkers for the diagnosis, prognosis, and treatment of several pathologies [7]. In recent years, our group has provided an overall picture of the EV proteome in several cardiovascular and cardiometabolic diseases. Initially, we identified novel potential EV-related biomarkers for ST-elevation myocardial infarction (STEMI) by comparing the proteome of EVs from STEMI patients to matched stable coronary artery disease (CAD) controls [8]. More recently, our group carried out a study comparing the proteome of circulating EVs from morbidly obese patients and lean individuals following two different proteomic approaches: 2D-DIGE-based and label-free MS-based proteomics. In that way, we identified a panel of biomarkers related to an increased cardiovascular risk in obesity [9]. The above approaches will be the basis for the protocols here presented. In this chapter, we will describe basic protocols of EV isolation and subsequent proteomic analysis. Moreover, we will focus on gel-based and quantitative MS-based approaches applied to the study of EVs in an attempt to identify novel EV biomarkers in a clinical context.

2 2.1

Materials Blood Collection

1. 3.2% sodium citrate tubes. 2. ACD: 78 mM citric acid, 96.6 mM trisodium citrate, and 111 mM glucose. 3. Eppendorf 5702 centrifuge and Eppendorf 5415 R.

2.2

EV Isolation

1. Beckman XL-100K Ultracentrifuge. 2. SW55 Ti rotor. 3. Ultra-clear 344057 tubes. 4. HEPES: 10 mM HEPES, 5 mM KCl, 1 mM MgCl2, 136 mM NaCl (pH 7.4). 5. 0.25 M KBr. 6. PBS: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4·12H2O, 2 mM KH2PO4, 1 L H2Omq (pH 7.4).

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15

7. DIGE buffer: 65 mM CHAPS, 5 M urea, 2 M thiourea, 0.15 M NDSB-256, 30 mM Tris Base, 1 mM sodium vanadate, 0.1 mM sodium fluoride, and 1 mM benzamidine. 8. LC-MS/MS buffer: 2% SDS, 500 mM Tris Base (pH 7.6), 0.05 M DTT. 2.3 Gel-Based Proteomics: 2D-DIGE

1. Coomassie plus protein reagent. 2. GE Healthcare CyDye™ DIGE Fluor Minimal Dye Labeling. 3. 10 mM lysine. 4. 2 sample buffer: 65 mM CHAPS, 2 M thiourea, 5 M urea, 0.15 M NDSB-256, 130 mM DTT, 4 mM tributylphosphine, 1 mM sodium vanadate, 0.1 mM sodium fluoride, and 1 mM benzamidine. 5. 4–7 Servalyt ampholytes. 6. 2D sample buffer: 5 M urea, 2 M thiourea, 2 mM tributylphosphine, 65 mM DTT, 65 mM CHAPS, 0.15 M NDSB-256, 1 mM sodium vanadate, 0.1 mM sodium fluoride, and 1 mM benzamidine. 7. pH 4–7 IPG, 24 cm strips. 8. Multiphor II system. 9. Reduction buffer: 6 M urea, 50 mM Tris Base (pH 8.8), 30% glycerol, 2% w/v SDS, 65 mM DTT, and traces of bromophenol blue. 10. Alkylation buffer: 6 M urea, 50 mM Tris Base (pH 8.8), 30% glycerol, 2% w/v SDS, 135 mM iodoacetamide, and traces of bromophenol blue. 11. 0.5% of melted agarose. 12. 11% polyacrylamide gels. 13. Ettan Dalt 6 system. 14. Typhoon FLA 7000 scanner. 15. Fixing buffer: 10% methanol/7% acetic acid. 16. SYPRO Ruby Protein Gel Stain. 17. Progenesis SameSpots software (v 4.5). 18. EASY-nLC Proxeon. 19. Bruker Amazon ETD ion trap. 20. Easy column SC200 Proxeon. 21. Data Analysis 4.0 and BioTools 3.2. 22. Mascot v2.3.0 database. 23. SwissProt human database.

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2.4 Gel-Free Proteomics: Label-Free

1. Pierce 660 nm Protein Assay, plus the Ionic Detergent Compatibility Reagent (IDCR). 2. 10 kDa Amicon™ Ultra-0.5. 3. 0.1 M NaOH. 4. UA buffer: 8 M Urea and 0.1 M Tris Base (pH 8.5). 5. 25 mM TEAB. 6. DTT 1 M in 25 mM TEAB. 7. 0.05 M iodoacetamide (IAA) in UA. 8. 0.1μg/μL trypsin in 20 mM TEAB. 9. Savant™ SPD121P SpeedVac. 10. 5% methanol/0.5% TFA. 11. Solvent A: 0.1% formic acid. 12. Solvent B: acetonitrile 0.1% formic acid. 13. Agilent 1200 nanoflow system. 14. C18 preconcentration cartridge. 15. C18 column. 16. LTQ-Orbitrap XL mass spectrometer. 17. Nanoelectrospray ion source. 18. Progenesis QI software (v 4.0). 19. Proteome Discoverer v1.4. 20. SwissProt human database.

3 3.1

Methods Blood Collection

The workflow of the experimental approach is shown in Fig. 1. Blood collection should be performed as recommended by the ISTH SSC collaborative workshop in order to minimize the impact of pre-analytical parameters on the measurement of circulating microparticles [10]. 1. Collect 50 mL of blood in coagulation 3.2% sodium citrate tubes (see Notes 1 and 2). 2. Add 450μL of warmed ACD to the blood and centrifuge it at 200  g for 20 min. 3. Pippete off the supernatant (platelet-rich plasma (PRP)) into a 50-mL centrifuge tube. 4. Add 10μg prostacyclin (10μL of stock solution at 1 mg/mL), mix gently by inversion, and immediately centrifuge at 1000  g for 10 min (see Note 3). 5. Pippete off the supernatant (platelet-poor plasma (PPP)) into a 50-mL centrifuge tube. Centrifuge it at 1500  g for 10 min.

Circulating EVs Clinical Proteomics

17

Fig. 1 Workflow of the experimental approach for the proteomic analysis of circulating EVs in a clinical context presented in this chapter

6. Transfer the supernatant into 1.5-mL tubes and centrifuge them at 15,000  g for 2 min using a fixed-angle centrifuge to obtain PFP (platelet-free plasma). 7. Freeze the samples at 3.2

EV Isolation

80  C until EV isolation (see Note 4).

Plasma EV isolation follows a protocol developed in our group based on the protocol initially established by Ramacciotti and colleagues [8, 9, 11]. 1. Add 1.2 mL of PFP per tube and mix it with 3 mL of HEPES buffer. In total, 7.4 mL of plasma is used per run. Ultracentrifuge them at 200,000  g for 90 min (4  C). 2. Remove the supernatant and resuspend the pellet with 4 mL of 0.25 M KBr. Incubate on ice for 20 min (see Note 5). 3. Spin down the samples for 90 min at 4  C, 200,000  g. 4. Remove the supernatant and wash the pellet with 4 mL of PBS in order to avoid plasma proteins. Centrifugate them under the same conditions.

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5. Dissolve the pellets depending on the proteomic approach: for 2D-DIGE, use 40μL of DIGE buffer; for label-free LC-MS/ MS, use 40μL of LC-MS/MS buffer. 6. Store EV lysates at

80  C.

3.3 EV Characterization

After isolation, there are several methods to characterize the heterogeneity of the EV sample. The ISEV recommends using at least two different technological approaches in order to characterize EV samples [2]. The principal techniques for characterization are electron microscopy-based (transmission electron microscopy (TEM) and cryo-TEM), particle enumeration techniques (e.g., nanoparticle tracking analysis (NTA), dynamic light scattering (DLS)), and flow cytometry [12].

3.4 Gel-Based Proteomics: 2D-DIGE Labeling

The example shown is based on a 2D-DIGE proteomic protocol where two different conditions are compared (e.g., health and disease). In this context, six gels were run as technical replicates using a total amount of 150μg of protein per gel. Due to the amount of protein required, pools can be used (see Note 6). All steps must be performed in the dark. 1. Label randomly 50μg of the protein mixture from each condition with 400 pmol minimal CyDye DIGE fluors Cy3 and Cy5. In addition, label 50μg of the third condition, which is a pool of both samples (internal standard) with 400 pmol Cy2. The labeling is on ice for 30 min. 2. Stop the reaction with 1μL of 10 mM lysine on ice for 10 min. 3. After the labeling step, pool the three samples labeled and add an equal volume of 2 sample buffer. Leave the mix for 15 min on ice. 4. Before reswelling of IPG strips, dilute the samples in 2D sample buffer to 500μL (final volume), and add ampholytes to a final concentration of 1.6% (v/v).

3.5 2D-Electrophoresis

1. Rehydrate the 24-cm 4–7 IPG strips with the sample for 16 h (see Note 7). 2. Run the first dimension (isoelectric focusing (IEF)) powered by the Multiphor II for 64.9 kVh at 17  C (see Note 8). 3. After the first dimension, equilibrate the strips with the reduction buffer for 15 min with gentle agitation. 4. Wash the excess with ultrapure water and, after that, equilibrate the strips with the reduction buffer for 15 min with gentle agitation. 5. Wash the strips again with ultrapure water. 6. Place the strips on the top of the second-dimension gels with 0.5% of melted agarose.

Circulating EVs Clinical Proteomics

19

7. Run the second dimension with SDS-polyacrylamide gel electrophoresis (PAGE) using 11% polyacrylamide gels. The running conditions are 20 mA per gel for 1 h and then 40 mA per gel for 4–6 h. All procedures are at 10  C using an Ettan Dalt 6 system. 8. Following the electrophoresis, scan directly the gels in a Typhoon FLA 7000 scanner. 9. After scanning, fix the gels in fixing buffer for 1 h and stain them with Sypro Ruby fluorescent dye for spot picking. 3.6 Differential Image Analysis

1. Perform the differential image analysis with Progenesis SameSpots software (v 4.5). 2. Align the images manually and automatically using the software. 3. Check the differentially regulated proteins present in the gels considering significant when the fold change is at least 1.5 and the p value is below 0.05. 4. Excise the differentially regulated spots manually from the gel for MS analysis.

3.7

MS Analysis

In the present example, samples are analyzed using LC-MS/MS on an EASY-nLC and a Bruker Amazon ETD ion trap. 1. Digest the spots in-gel with trypsin using a protocol defined by Shevchenko et al. with minor modifications [13]. 2. Separate the digested peptide mixtures dissolved in 0.1% formic acid in EASY-nLC with a reverse-phase nanocolumn (Easy column SC200). 3. Use a CID-ETD ion trap mass spectrometer equipped with a Nanosprayer ionization source to analyze the ionized peptides. The spectra were acquired in Enhanced Resolution mode. 4. Analyze the mass data with Data Analysis 4.0 and BioTools 3.2. Search the results with the database Mascot v2.3.0 search tool screening SwissProt (see Note 9). 5. Accept as positive only these identifications that obtain more than 50% y-ions (CID fragmentation) or z-ions (ETD fragmentation) for peptide comprising at least eight amino acids long and no missed tryptic cleavage site. Positive hits correspond to Mascot scores >40 plus the fulfillment of the above criteria.

3.8 Gel-Free Proteomics: Label-Free

Another approach that can be used to compare different EVs from different sources (e.g., health and disease) is label-free LC-MS/ MS. In this case, protein samples were digested with sequencing grade modified trypsin using FASP (filter aided sample preparation protocol). All steps are run at 13  C.

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Maria N. Barrachina and A´ngel Garcı´a

FASP Digestion

1. Before digestion, prepare the filters of the Amicon™ Ultra-0.5. Wash the membranes with 400μL of 0.1 M NaOH at 14,000  g for 10 min. Add another wash with 400μL ultrapure water and centrifugate it. 2. Prepare the samples: thaw them for 10 min using Eppendorf Thermomixer® comfort at 600 rpm and 20  C. 3. Add 0.1 M DTT in order to obtain 20 mM (final concentration). Heat samples for 5 min at 99  C in the Thermomixer. 4. Temper the samples before adding UA buffer. Dilute samples with UA buffer to obtain less than 0.1% SDS (final concentration) (see Notes 10 and 11). 5. Add the samples into the filter and then centrifuge for 15 min at 14,000  g. The maximum volume inside the filter is 450μL. Do as many steps as needed in order to introduce all samples into the filter. 6. Wash the filter two times with 200μL UA buffer and centrifuge them for 15 min at 14,000  g. 7. Alkylation step: add 0.05 M IAA for 20 min with gentle agitation (400 rpm). 8. Wash two times the filter with 200μL UA buffer and then four times with TEAB buffer. All steps are followed with a centrifugation for 15 min at 14,000  g (see Note 12). 9. Digest with trypsin (5:100) (v/v) for 18 h with gentle agitation at 37  C in the dark. 10. The second day, elute the samples with 100μL TEAB (threetimes) and centrifuge for 15 min at 14,000  g. 11. Evaporate the elute with a SpeedVac and resuspend the digested peptides in 20μL of 5% methanol; 0.5% TFA.

3.10 LC-MS/MS Analysis

In this example, samples are analyzed using LC coupled to an LTQ XL Orbitrap equipped with a nanoESI ion source. 1. Dilute the sample to a final volume of 20μL and load it into the chromatographic system consisting of a C18 preconcentration connected to a 15-cm-long, 100-μm-i.d. C18 column. 2. Separate them using 0.4μL/min in a 120-min acetonitrile gradient from 3% to 35% of solvent A using an Agilent 1200 nanoflow system (see Note 13). 3. The LC is coupled to an LTQ-Orbitrap XL mass spectrometer equipped with a nanoelectrospray ion source (see Note 14). 4. Use the database search Proteome Discoverer v1.4 and SwissProt human database in order to identify proteins.

Circulating EVs Clinical Proteomics

3.11 Differential Analysis for Label-Free

21

1. Use Progenesis QI software (v 4.0). Do not filter the peptides’ features. 2. Obtain the mascot generic file (.mgf) generated from all exported MS/MS spectra. Use SEQUEST (Proteome Discoverer v1.4) with the following parameters: peptide mass tolerance 20 ppm, fragment tolerance 0.6 Da, enzyme set as trypsin and allowance up to one missed cleavages, dynamic modification of methionine oxidation (+16 Da), and fixed modification of cysteine carbamidomethylation (+57 Da). 3. Filter the results with peptide rank 1 and peptide confidence high (1% FDR). 4. After that, import this file into Progenesis QI software for further analysis. The statistic criteria are ANOVA test with p value less than 0.05 and fold changed 1.5.

3.12

4

System Biology

In order to obtain more information about the proteins identified, several open-access software can be used. For example, Cytoscape and Funrich software can be used to investigate possible interactions among all the identified proteins and their molecular functions. In particular, Cytoscape is an open-source software platform, which is used to visualize complex networks and integrate these with any type of attribute data [14]. Moreover, it is also a complementary tool to know the biological process, molecular functions, and cellular components of the differential proteins. In addition, FunRich software shows the cellular component, the biological process, and the biological pathways among all proteins identified [15]. Furthermore, there are different EV-related databases that can be used, such as EVpedia and Vesiclepedia, to confirm that the proteins identified were enriched in EVs. Finally, the EV biomarkers identified must be validated in an independent cohort of patients bay target MS or other western blotting-based approaches (e.g., enzyme-linked immunosorbent assay (ELISA) or western blot).

Notes 1. Plasma is usually the preferred source of EVs because additional EVs are released during clot formation when preparing serum. 2. The SSC on Vascular Biology of ISTH proposes that EV-containing plasma must be isolated within 2 h after blood collection. 3. Prostacyclin avoids unwanted platelet activation during centrifugation by raising the intracellular concentration of cAMP. This prevents the release of EVs from platelets during centrifugation.

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Maria N. Barrachina and A´ngel Garcı´a

4. The recommendation is a single freeze–thaw cycle and storage up to 1 year ( 80  C) since these conditions have a low impact on the concentration and size of vesicles. 5. KBr is used to solubilize lipoproteins and remove them from the plasma. 6. The use of pools is not ideal but can be the only choice due when there is sample limitation. In that case, validation should be in individual biological samples. 7. IPG strips pH 4–7 are preferred for use because the majority of EV proteins are distributed in this pH range. 8. If other equipment is used (e.g., IPGPhor 3), the electrophoresis protocol must be adapted. 9. Searches are restricted to human taxonomy allowing carbamidomethyl cysteine as a fixed modification and oxidized methionine as potential variable modification. Both the precursor mass tolerance and the MS/MS tolerance were set at 0.3 and 0.4 Da, respectively, allowing one missed tryptic cleavage site. 10. With increasing temperature, urea decomposes to ammonia and isocyanic acid. For this reason, it is necessary to temper samples before diluting them with UA buffer. 11. Only less than 0.1% SDS (final concentration) is compatible with the filter. For this reason, it is necessary to dilute the sample many times with the UA buffer in order to reduce the concentration. 12. The trypsin activity is affected by the concentration of urea. Therefore, mix gently the washes with the pipette in order to avoid possible remains of urea. 13. The HPLC system mentioned in this example is composed of an Agilent 1200 capillary nano pump, a binary pump, a thermostated microinjector, and a micro switch valve. 14. The LTQ XL Orbitrap is operated in the positive ion mode with a spray voltage of 1.8 kV. The spectrometric analysis is performed in a data-dependent mode, acquiring a full scan followed by ten MS/MS scans of the ten most intense signals detected in the MS scan from the global list. The full MS (range 400–1650) is acquired in the Orbitrap with a resolution of 60,000. The MS/MS spectra are done in the linear ion trap.

Acknowledgments The authors acknowledge support from the Spanish Ministry of Science and Innovation [grants No. SAF2016-79662-R, and PID2019-108727RB-I00], co-funded by the European Regional Development Fund (ERDF). Financial support from the

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Consellerı´a de Cultura, Educacio´n e Ordenacio´n Universitaria, Xunta de Galicia (Centro Singular de investigacio´n de Galicia accreditation 2019–2022, ED431G 2019/02), and the European Regional Development Fund (ERDF) is also acknowledged. References 1. Yuana Y, Sturk A, Nieuwland R (2013) Extracellular vesicles in physiological and pathological conditions. Blood Rev 27:31–39 2. Lo¨tvall J, Hill AF, Hochberg F, Buza´s EI, Di Vizio D, Gardiner C, Gho YS, Kurochkin IV, Mathivanan S, Quesenberry P, Susmita S, Tahara H, Wauben MH, Witwer KW, The´ry C (2014) Minimal experimental requirements for definition of extracellular vesicles and their functions: a position statement from the International Society for Extracellular Vesicles. J Extracell Vesicles 3:26913 3. Heijnen HF, Schiel AE, Fijnheer R, Geuze HJ, Sixma JJ (1999) Activated platelets release two types of membrane vesicles: microvesicles by surface shedding and exosomes derived from exocytosis of multivesicular bodies and alphagranules. Blood 94:3791–3799 4. Simons M, Raposo G (2009) Exosomes— vesicular carriers for intercellular communication. Curr Opin Cell Biol 21:575–581 5. Colombo M, Moita C, van Niel G, Kowal J, Vigneron J, Benaroch P, Manel N, Moita LF, The´ry C, Raposo G (2013) Analysis of ESCRT functions in exosome biogenesis, composition and secretion highlights the heterogeneity of extracellular vesicles. J Cell Sci 126:5553–5565 6. Abels ER, Breakefield XO (2016) Introduction to extracellular vesicles: biogenesis, RNA cargo selection, content, release, and uptake. Cell Mol Neurobiol 36:301–312 7. Gaceb A, Martinez MC, Andriantsitohaina R (2014) Extracellular vesicles: new players in cardiovascular diseases. Int J Biochem Cell Biol 50:24–28 ˜a AF, Ocaranza-Sa´nchez R, 8. Ve´lez P, Parguin Grigorian-Shamagian L, Rosa I, AlonsoOrgaz S, de la Cuesta F, Guitia´n E, Moreu J, ´ Barderas MG, Gonza´lez-Juanatey JR, Garcı´a A (2014) Identification of a circulating microvesicle protein network involved in ST-elevation myocardial infarction. Thromb Haemost 112:716–726 9. Barrachina MN, Sueiro AM, Casas V, Izquierdo I, Hermida-Nogueira L, Guitia´n E, Casanueva FF, Abia´n J, Carrascal M, Pardo M, ´ (2019) A combination of proteomic Garcı´a A approaches identifies a panel of circulating extracellular vesicle proteins related to the risk

of suffering cardiovascular disease in obese patients. Proteomics 19:e1800248 10. Vila-Liante V, Sa´nchez-Lo´pez V, Martı´nez˜ ez LA, Arellano-Orden E, Sales V, Ramo´n-Nun Cano-Ruiz A, Rodrı´guez-Martorell FJ, Gao L, Otero-Candelera R (2016) Impact of sample processing on the measurement of circulating microparticles: storage and centrifugation parameters. Clin Chem Lab Med 54:1759–1767 11. Ramacciotti E, Hawley AE, Wrobleski SK, Myers DD, Strahler JR, Andrews PC, Guire KE, Henke PK, Wakefield TW (2010) Proteomics of microparticles after deep venous thrombosis. Thromb Res 125:e269–e274 12. Barrachina MN, Caldero´n-Cruz B, Fernandez´ (2019) Application of extraRocca L, Garcı´a A cellular vesicles proteomics to cardiovascular disease: guidelines, data analysis, and future perspectives. Proteomics 19:e1800247 13. Shevchenko A, Jensen ON, Podtelejnikov AV, Sagliocco F, Wilm M, Vorm O, Mortensen P, Shevchenko A, Boucherie H, Mann M (1996) Linking genome and proteome by mass spectrometry: large-scale identification of yeast proteins from two dimensional gels. Proc Natl Acad Sci U S A 93:14440–14445 14. Shannon P, Markiel A, Ozier O, Baliga NS, Wang JT, Ramage D, Amin N, Schwikowski B, Ideker T (2003) Cytoscape: a software environment for integrated models of biomolecular interaction networks. Genome Res 13:2498–2504 15. Pathan M, Keerthikumar S, Chisanga D, Alessandro R, Ang CS, Askenase P, Batagov AO, Benito-Martin A, Camussi G, Clayton A, Collino F, Di Vizio D, Farcon-Perez JM, Fonseca P, Fonseka P, Fontana S, Gho YS, Hendrix A, Hoen EN, Iraci N, Kastaniegaard K, Kislinger T, Kowal J, Kurochkin IV, Leonardi T, Liang Y, Llorente A, Lunavat TR, Maji S, Monteleone F, Øverbye A, Panaretakis T, Patel T, Peinado H, Pluchino S, Principe S, Ronquist G, Royo F, Sahoo S, Spinelli C, Stensballe A, The´ry C, van Herwijnen MJC, Wauben M, Welton JL, Zhao K, Mathivanan S (2017) A novel community driven software for functional enrichment analysis of extracellular vesicles data. J Extracell Vesicles 6:1321455

Chapter 3 Utilization of Laser Capture Microdissection Coupled to Mass Spectrometry to Uncover the Proteome of Cellular Protrusions Ana Gordon and Karine Gousset Abstract Laser capture microdissection (LCM) provides a fast, specific, and versatile method to isolate and enrich cells in mixed populations and/or subcellular structures, for further proteomic study. Furthermore, mass spectrometry (MS) can quickly and accurately generate differential protein expression profiles from small amounts of samples. Although cellular protrusions—such as tunneling nanotubes, filopodia, growth cones, invadopodia, etc.—are involved in essential physiological and pathological actions such as phagocytosis or cancer-cell invasion, the study of their protein composition is progressing slowly due to their fragility and transient nature. The method described herein, combining LCM and MS, has been designed to identify the proteome of different cellular protrusions. First, cells are fixed with a novel fixative method to preserve the cellular protrusions, which are isolated by LCM. Next, the extraction of proteins from the enriched sample is optimized to de-crosslink the fixative agent to improve the identification of proteins by MS. The efficient protein recovery and high sample quality of this method enable the protein profiling of these small and diverse subcellular structures. Key words Cellular protrusion, Laser capture microdissection, Proteomics, Fixation, DTBP, Mass spectrometry

1

Introduction Laser capture microdissection (LCM) is a tool that facilitates, via direct visualization of cells or tissues, the enrichment and isolation of cells of interest in a heterogeneous sample [1]. There are different classes of LCM depending on the type of laser and capture method. Overall, laser capture microdissection is an optical microscope, which can be coupled to fluorescence, with a finely focused laser and a system for capturing dissected samples. The cells of interest can be selected from the heterogeneous sample through a touch screen, enabling the selection of regions of interest (ROI) using an interactive pen or the mouse. Each ROI can be drawn freehand or using predefined geometrical shapes. This technique

Mo´nica Carrera and Jesu´s Mateos (eds.), Shotgun Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 2259, https://doi.org/10.1007/978-1-0716-1178-4_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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has been successfully applied in the isolation of tumoral cells, neurons affected by Parkinson or Alzheimer diseases, virus-infected cells, etc. [2–4]. Moreover, LCM was shown to be able to separate cellular organelles from specific cells [5], growth cones [6], and invadopodia [7]. One advantage of using LCM to enrich a sample with the cells of interest is the possibility of performing subsequent downstream studies of DNA, RNA, or protein composition. Proteomics, the study of proteins, is one of the most important technologies used to acquire new insights into cellular function since proteins are the primary effectors of all biological activity. However, the feasibility of combining proteomic techniques with LCM depends primarily on the sensitivity of the technique used, as the amount of protein retrieved from the dissected sample can be very small. Traditional proteomic techniques cannot solve this problem since they require a high degree of sample homogeneity to obtain significant results. Nevertheless, mass spectrometry can be applied to identify and quantify proteins from different samples. In microproteomics, much smaller amounts, in the attomole or even zeptomole range, are required to sequence a peptide [8]. Cellular protrusions are involved in several biological functions, such as cell migration, neurite outgrowth, phagocytosis, and cellto-cell communication. More recently, these structures have also been related to cancer-cell invasion, intercellular transmission of misfolded proteins in degenerative diseases, and intercellular spread of infectious pathogens. These physiological and pathological processes are conducted by different types of cellular protrusions, such as lamellipodia, pseudopodia, filopodia, growth cone, tunneling nanotubes, and others [9–17]. However, there are many unanswered questions related to their mechanism of formation, their structural components, or signaling pathways [18]. One of the principal reasons for the lack of knowledge regarding cellular protrusions is the limited techniques that allow studying them specifically. Previous attempts have been made to isolate cellular protrusions by using a Boyden chamber [19–21] or an excimer laser [22]. However, none of these methods are specific since they do not visualize what they are isolating and they were only able to enrich pseudopodia, a specific subtype of cellular protrusion, but could not specifically isolate different types of cellular protrusions. Here, we describe a protocol that combines two techniques to enhance our knowledge of the composition of these structures: LCM, to get an enrichment of cellular protrusions based on morphological features, and MS, to uncover the proteomes of these structures. The first roadblock in using LCM to isolate cellular protrusions is their transient nature and structural fragility. Thus, we first needed to identify fixative procedures that would be strong enough to maintain the structural integrity of the cellular protrusions for laser microdissection but also compatible with downstream

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27

microproteomic analyses. Fixative solutions with glutaraldehyde are ideal for maintaining cellular protrusions, including fragile protrusions such as tunneling nanotubes [23]. Unfortunately, glutaraldehyde fixations, even in low concentrations, create thorough cross-linking that seriously hampers the quality sample for MS analysis [24]. Here, we use dimethyl-3-30 -dithiobispropiomimidate (DTBP), which can be de-cross-linked by using a reducing agent and a temperature-step process [25, 26]. In fact, the currently optimized fixation with DTBP not only preserves cellular protrusions but also allows for the identification of high-quality and highquantity proteins, which reduces the number of cellular protrusions needed for MS analysis [27]. The current protocol allows one to specifically select any type of cellular protrusion based on morphological and/or fluorescence markers, thus greatly improving its selectivity compared to previous methods [6, 7, 19–22]. Importantly, we were able to isolate TNTs, a very delicate and specialized cell-to-cell communication conduit, which have been related to several diseases, such as cancer, human immunodeficiency virus, or prion infection [28–31].

2 2.1

Materials Cell Culture

1. CAD (cath-a-differentiated) cell line. 2. 25-cm2 cell culture flasks. 3. 5-, 10-, and 25-mL individually wrapped pipettes. 4. 1-mL aspirating pipettes. 5. 15- and 50-mL conical tubes. 6. An optical biological inverted microscope. 7. A biological safety cabinet, e.g., 1300 Series Class II, Type A2 Biological Safety Cabinet. 8. CO2 incubator and an aspiration pump. 9. CAD cell culture medium: Add 50 mL of 10% fetal bovine serum (FBS) to 450 mL of OPTI-MEM. Filter-sterilize and store at 4  C for no longer than 4 weeks. 10. 0.4% trypan blue solution. 11. Metalized hemocytometer.

2.2 Preparation for LCM

As stated above, there are different types of LCM. In our studies, we used a laser microdissection system controlled by the Molecular Machines & Industries (MMI) Cell Tools Software. 1. MMI live cell chamber with membrane and petri dish. This includes a membrane ring for the initial cultivation of cells and a UV-permeable microdissection chamber to isolate the cellular protrusions of interest.

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2. 10 phosphate-buffered saline (PBS), pH 7.4 and fibronectin solution from bovine plasma. 3. Tweezers, filter tips, pipettes, and an aspiration pump. 2.3

Fixation

1. 30% hydrogen peroxide (H2O2). 2. 16% paraformaldehyde (PFA), EM grade. 3. 1 M HEPES. 4. pH meter. 5. Vortex mixer and chemical hood. 6. Microcentrifuge tubes. 7. Pipette tips and Pipetman classic™ Starter Kit. 8. Cell membrane marker (wheat germ agglutinin tetramethylrhodamine conjugate). 9. Ibidi μ-dishes. 10. Fluorescence microscope. 11. Fixation solution 1: 4% PFA solution, 200 mM HEPES, and 1 PBS (see Note 1). 12. Fixation solution 2: 5 mM DTBP solution. Weigh DTBP powder to make up a 50 mM DTBP stock and dissolve in 1 PBS, pH 8.6. Calculate the necessary volume of the stock solution to get a final concentration of 5 mM DTBP and add 25 mM HEPES in 1 PBS, pH 8.6 (see Note 2).

2.4 Laser Capture Microdissection 2.5 Protein Extraction

Molecular Machines & Industries (MMI) CellCut Laser Microdissection and the Software (MMI Cell Tools Software; Ver. 4.3.3.). 1. Digital dry bath (37, 60, and 100  C). 2. Sonicator. 3. Refrigerated microcentrifuge. 4. Bovine serum albumin standard. 5. RC DC™ Protein Assay Kit II. 6. Truview cuvettes. 7. A spectrophotometer, Spectrophotometer.

e.g.,

Smartspec™

Plus

8. Alternatively, samples can be acquired based on the number of cellular protrusions isolated by LCM, rather than protein assay. 9. Lysis buffer (RIPA): 10 mM Tris–HCl (pH 8.0), 1 mM EDTA, 0.5 mM EGTA, 2% SDS, 1% Triton X-100, 0.1% sodium deoxycholate, 140 mM NaCl, 100 mM DTT (see Note 3), 1% protease inhibitors (e.g., Halt™ Protease Inhibitor Cocktail, 100), and molecular biology grade water.

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2.6 Mass Spectrometry Sample Preparation

29

1. 5 Laemmli buffer: 25 mM Tris–HCl (pH 6.8), 25% glycerol, 10% SDS, 0.02% bromophenol blue, 150 mM DTT, and molecular water (see Note 4). 2. An 8% acrylamide Bis-Tris gel for mass spectrometry: Prepare a 1.25 M Bis-Tris stock and adjust pH to 6.5–6.8. For the resolving gel, calculate the necessary volume of the stock solution to get a final concentration of 350 mM Bis-Tris and add the corresponding volume of 30% acrylamide/bis-acrylamide (37.5:1) to get a final concentration of 8%. Add H2O to make up a final volume of 5 mL. Finally, add 10% ammonium persulfate (APS) and TEMED (10 μL) to polymerize it. For the stacking gel, calculate the necessary volume of the stock solution to get a final concentration of 350 mM Bis-Tris and add the corresponding volume of 30% acrylamide to get a final concentration of 4%. Add H2O to make up a final volume of 1.75 mL. Finally, add 10% APS and TEMED (10 μL) to polymerize it. 3. 20 MOPS buffer: 1 M 3-(N-morpholino) propane sulfonic acid (MOPS), 1 M Tris Base, 20 mM EDTA, 2% SDS, and molecular water (see Note 5). 4. Running buffer: 20% MOPS buffer, 0.5% sodium bisulfite, and molecular water (see Note 1). 5. Fixing solution for gels: methanol, acetic acid, and molecular water (40:40:8, respectively) (see Note 1). 6. Precision Plus Protein™ Dual Color Standards (Bio-Rad, #161-0374), loading tips. 7. Powerpac™ Basic Power Supply and Mini-PROTEAN® Tetra cell system. 8. Compact digital rocker. 9. Speed vacuum. 10. iST Sample Preparation Kit from PreOmics.

2.7 Mass Spectrometers

3 3.1

1. Thermo Scientific Orbitrap Fusion mass spectrometer or timsTOF Pro mass spectrometer from Bruker.

Methods Cell Culture

The method described here is to get an enrichment of cellular protrusions and, subsequently, to extract the proteins and analyze them by MS. Cell density is a critical step when isolating cellular protrusions. It can vary not only with the types of cellular protrusions that need to be isolated but also with the cell types used for the experiments. For example, if you are interested in isolating filopodia, we recommend plating low cell density in order to ensure

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that filopodia can be identified and selected from single cells. If you are interested in isolating TNTs, then the cells need to be in closer proximity in order to be able to form TNTs and therefore a higher cell density is required. These numbers must be predetermined for each cell types used, as well as for the types of cellular protrusion of interest. 1. Pre-warm a 50-mL centrifuge tube with CAD media in the 37  C water bath. Place 3 mL of media into a 15-mL conical tube and 5 mL of media into a T25 plate. Remove the cells from the liquid nitrogen tank. Thaw the cryotube with CAD cells in the 37  C water bath. When almost thawed (it is acceptable for some ice crystals to be left), take the tube out of the water bath and move to the tissue culture hood. As freezing media contains DMSO, which is toxic for cells, these steps have to be done quickly. 2. Add 1 mL of media to the cells (approximately 1,500,000 cells), resuspend them, and transfer them into the prepared 15-mL conical tube. Spin the conical tube in a centrifuge at 1000  g for 4 min (the cells will collect at the bottom as a pellet). Carefully aspirate and discard the supernatant from the 15-mL conical tube and add 1 mL of media to the cell pellet and resuspend gently. Then, pull up all of the cell mixture into the pipette and transfer it to the prepared T25 plate. Look at cells before placing them in the incubator. The cells should be round and shiny. 3. Next day, check that the cells have adhered to the plate and are healthy and nondifferentiated and look at their confluency. Grow cells until they become 80% confluent (usually 2 days). 4. To split the cells, remove the media and dead cells, wash once with PBS, and add 1 mL of media. Tap the flask to detach the cells (mechanical dissociation) and observe under the microscope to ensure that the majority of the cells are detached. Add 3 mL of fresh media along the walls to scrape out all the cells. Pipette out the total 5 mL of the cultures, by pressing the tip of the pipette on the bottom of the flask to break clumps and check under the microscope to ensure that you have mostly single cells. Add 1 mL of the culture (approximately 1,000,000 cells) to the new flask and take the volume of fresh media needed to make up the new flask to 6 mL. 5. Alternatively, cells can be grown in a 60- or 10-cm culture dish. In this case, cells can be detached mechanically by pipetting them up and down, rather than tapping the flask. 6. After 2 days, cells should be 80% confluent, and they can be split again. Cells should be split a minimum of two times before seeding them in an MMI live chamber dish to ensure cell viability. Once the cells have been split two times, they can be

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counted by mixing a sample of the cell solution with Trypan blue (dilution 1:1). Count the cells using a hemocytometer or an automated cell counter. Calculate the volume for your desired cell concentration (i.e., 200,000 cells for TNTs) to be seeded in an MMI live chamber dish. 3.2 Differentiated CAD Cells

In order to obtain axonal and dendritic protrusions, we differentiated CAD cells by serum starvation. 1. Seed 30,000 CAD cells in an MMI live chamber dish. 2. To differentiate these cells, they have to be grown in OPTIMEM without FBS for 10 days. 3. The rest of the protocol is the same as for CAD cells.

3.3 Preparation of MMI Live Chamber Dish

Optimal conditions for cell attachment will depend on the cell type and should be predetermined. For CAD cells, we use fibronectin coating, but there are other extracellular matrix protein coatings available for different cell lines (see Note 6). 1. Remove the membrane ring from the 35-mm dish with regular tweezers and transfer it to the adhesive area of the UV-permeable microdissection chamber. Press carefully with your fingers and, once attached, do not move the ring to prevent wrinkles and/or breakages (see Note 7). 2. To confirm that the membrane ring is properly attached and does not leak, add 1 mL of filtered PBS to the dish and wait for at least 5 min. If no leaks are observed, proceed with the sample preparation. If PBS is seen on the outside of the membrane ring, discard the ring/chamber and start anew. 3. Prepare 300 μL of fibronectin (6 mM).

filtered

PBS

and

24

μL

of

4. Remove the PBS, add the fibronectin solution, and make sure that the fibronectin solution covers the entire surface of the ring since the membrane is hydrophobic. Place the dish in the 37  C/5% CO2 incubator for 20 min. 5. Wash three times with filtered PBS and two times with CAD media. 6. Add the desired volume of cells (in our case, 200,000 cells for TNT samples) and leave them in the incubator until ready for fixation (see Note 8). 7. Several protocols can be applied to increase cellular protrusions and should be predetermined based on your cell types and cellular protrusion of interest. We have included two of them (serum starvation for axons/dendrites or exposure to H2O2 for TNTs), but many other stimuli can be applied depending on your research goals (see Note 9).

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Ana Gordon and Karine Gousset

Fixation

As previously stated, cellular protrusions are fragile and must be fixed prior to LCM isolation [26, 27]. However, while a strong fixative like glutaraldehyde is ideal for maintaining the integrity of cellular protrusions, it is irreversible and does not allow for efficient protein extraction post cell lysis (Fig. 1a, b). On the other hand, we have recently demonstrated that using a cleavable crosslinker like DTBP not only preserves cellular protrusions [26] but it also allows for a greatly improved protein recovery after cell lysis (Fig. 1c). Overall, the use of DTBP along with the alternate protein extraction protocol we have developed [26] allows for the reduction of the overall number of cellular protrusions needed to be isolated by LCM, as it greatly increases the number of spectra, number of proteins, and number of total unique proteins identified by MS compared to glutaraldehyde-fixed samples (Fig. 2). 1. To increase the number of cellular protrusions, a concentration of 100 μM of H2O2 in warm CAD media is added to the cells for 5 min (see Note 10). 2. Add 100 μL for 500 μL total volume or 200 μL for 1 mL total volume of the fixation solution 1 (4% PFA solution) directly to the cells in their culture media and wait for 5 min. 3. Aspirate and add 1 mL of fixation solution 1 for 15 min (see Note 11). Change fixative solutions by tilting the dish, allowing and carefully aspirating the solution near the wall, without disturbing cells, or they might detach and cellular protrusions could break. Take the same considerations for steps 4 and 5. 4. Aspirate and add 1 mL of fixation solution 2 (5 mM DTBP solution) for 15 min. 5. Aspirate and add 1 mL of filtered PBS (see Note 12). We recommend preestablishing the best conditions (time and concentration) for induction of cellular protrusions with H2O2 for different cell types. Similarly, different concentrations of DTBP might be required to optimize the maintenance of cellular protrusions for different cell types. Trials should be carried out with a cell membrane marker (e.g., wheat germ agglutinin tetramethylrhodamine conjugate (WGA-Rhod, 1:200 in PBS) in Ibidi μ-dishes (Ibidi)) and check for cellular protrusion stability under a fluorescence microscope [26]. The cells should be washed twice with PBS after fixation and labeled for 10–20 min in the dark at room temperature with WGA-Rhod.

3.5 Laser Capture Microdissection (LCM)

LCM parameters have to be set up to improve the isolation of different cellular protrusions from the cells of interest. Here are some key steps to take into account when using the LCM. 1. Mount the microdissection chamber dish onto the stage of the LCM microscope and open the MMI Cell Tool program.

Proteome of Cellular Protrusions Identified by LCM/MS

33

Fig. 1 Effect of fixations, protein extractions, and dehydration of samples on protein yield. GLU/PFA fixed samples (a) lysed with RIPA buffer (0.01% SDS) on ice or (b) lysed with RIPA buffer (2% SDS) and incubations at 100  C/20 min and 60  C/2 h; (1) unfixed lysates, (2) 4% PFA alone, (3) GLU (0.05%)/PFA, (4) GLU (0.01%)/ PFA, and (5) GLU (0.005%)/PFA; (c) PFA/DTBP fixed samples were lysed with RIPA buffer (2% SDS and 100 mM DTT) and incubations at 37  C/30 min, 100  C/20 min, and 60  C/2 h; (1) unfixed lysates, (2) 4% PFA alone, (3) PFA/3 mM DTBP, (4) PFA/5 mM DTBP, and (5) PFA/10 mM DTBP. (d) PFA/5 mM DTBP fixed samples extracted and loaded as in gel C; (1) cells were fixed and lysed the same day; (2) cells were fixed and kept in PBS for 24 h before lysing; (3) cells were fixed and kept dry for 24 h before lysing; (4) cells were fixed and kept in PBS for 72 h before lysing; and (5) cells were fixed and kept dry for 72 h before lysing. Stars indicate protein bands with decreased intensity. Gels are representative of three independent experiments. (Reproduced from ref. 26 with permission from Proteomics)

2. The CellCut mode of the software allows for the calibration of the slide geometry of the dish that keeps the plane of focus steady as the laser cuts around the protrusions. With the 4 objective, focus the dish and set the proper plane tilt of the dish. Then, scan the dish to visualize the entire cell-covered area and help in determining what area to cut.

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Fig. 2 Effect of type of fixation on protein yield and MS results from 1000 cells. (a) Silver-stained representative gel from (1) unfixed cells (0.1 μg of protein); (2) 1000 GLU/PFA fixed cells; and (3) 1000 PFA/DTBP fixed cells; (b) average and standard deviation of unique proteins, unique spectra, and unique peptides of samples from 1000 PFA/DTBP or GLU/PFA fixed cells. (Reproduced from ref. 26 with permission from Proteomics)

3. The MMI cell tools software also allows the manipulation of the laser parameters (speed, focus, and power) to get a precise cut of the desired protrusions. The settings for LCM have to be adjusted for each sample with the objective required to cut the cellular protrusions of interest. Different objectives may be used depending on the cellular protrusions of interest (see Fig. 3). 4. The laser speed should be slowed down to avoid detaching the cells or disturbing the cellular protrusions during the process of cutting. This will need to be determined for each sample depending on the types of isolated cellular protrusions. For example, for growth cones or filopodia, which are strongly attached to the substratum, the cut velocity can be higher (e.g., 35 μm/s). On the other hand, for more fragile structures like TNTs, the cut velocity should be reduced (e.g., 10 μm/s). 5. Laser focus is a way to adjust the position of the laser beam in the Z-direction within the sample, determining the thickness of the cut. It should neither be too low to detach the membrane as it is cutting, nor too high to not cut the membrane at all. This will be determined by calibrating the laser focus according to the plane tilt of the dish. Thus, the laser focus should be adjusted for each experiment as it may vary from dish to dish and sample to sample. 6. The power needed to cut the sample is proportional to the sample thickness. It is represented in the percentage of UV light transmitted by the laser. The laser power should be set

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Fig. 3 Cellular protrusions isolated by laser capture microdissection. LCM, which uses a finely focused laser to cut a region of interest (ROI), is ideal for isolating cellular protrusions. Cells were plated on MMI live cell chambers, fixed, and

Ana Gordon and Karine Gousset

to the minimal value sufficient to cut through the membrane, and we suggest having at least two laser cut repetitions (i.e., the laser will cut the same area twice). This is critical to ensure that the membrane is properly cut from the rest of the samples since these are the samples of interest that will need to stay attached to the chamber after removal of the ring (in our case, 75–85% of laser power was enough). 7. Using the proper objective (depending on the size of the cellular protrusions of interest) [27], you can select the ROIs by manually drawing a line with a digital pen, directly on the computer screen, around the protrusions of interest (Fig. 3 shows different types of cellular protrusions that can be isolated by LCM). 8. We recommend selecting 10–100 ROIs at a time, prior to cutting, but these can be changed according to need or user preferences. 9. Between cutting sessions, replace the PBS with fresh PBS to remove any dead cells or debris and store the dish at 4  C until further use. 10. The dish can be used for 4–5 days approximately. Thus, as many ROIs should be isolated as possible per day and a new dish needs to be prepared if more ROIs are required after 5 days post-fixation (see Note 13).

ä

36

Fig. 3 (continued) imaged at 20 (a), 40 (b–d), or 60 (e) magnifications. Various types of cellular protrusions are shown. For all cases, (i) ROIs are drawn around the protrusions of interest, (ii) are representative images of cellular protrusions after the laser cut, and (iii) are images of the desired isolated cellular protrusions after removal of the LCM membrane containing the “unwanted” cells. Examples of different types of protrusions are shown: (a) axons and dendrites from dCADs are shown (scale bar ¼ 100 μm). Small dendritic filopodia can be observed (black arrows); (b) in order to increase the number of cellular protrusions, CAD cells were treated for 5 min with 100 μM H2O2 prior to fixation. Various types of cellular protrusions are shown (scale bar ¼ 10 μm). Individual subtypes of cellular protrusions can be specifically isolated such as (c) GCs (scale bar ¼ 20 μm), (d) filopodia (scale bar ¼ 10 μm), and (e) TNTs (scale bar top ¼ 50 μm; bottom ¼ 10 μm). TNTs do not touch the substratum, and tension is visible within these structures (ei). As expected, after being cut, the structures collapsed onto the LCM membrane (eii,iii), clearly demonstrating that these protrusions were TNTs, and not attached filopodia or other types of protrusions. (Reproduced from ref. 27 with permission from International Journal of Molecular Sciences)

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11. To isolate the ROIs cut, remove the PBS, wash the dish gently with filtered PBS, aspirate the PBS, and carefully lift the ring from the microdissection chamber with tweezers. All of the desired ROIs should be left on the sticky membrane of the microdissection chamber, while the rest of the cells will be removed along with the ring. It is imperative to go back to the LCM and look at the isolated ROIs left on the microdissection chamber to ensure that the expected ROIs are present and that no whole cells or other cellular debris are present. If a cell is found, it must be obliterated with the laser. 12. Once the ROIs have been examined and validated, the cellular protrusions can be lifted from the microdissection chamber with a micropipette by taking a small volume of lysis buffer (i.e., 5 μL) and pipetting up and down all over the microdissection chamber where the cuts are located (see Note 14). Avoid making bubbles. The cuts should be transferred to a 1.5-mL centrifuge tube, and the process can be repeated to ensure that all cuts have been lifted by observing the dish under the microscope. The final volume should be small (i.e., 15 μL total for all of your samples). 13. For the whole cell lysate sample, cut the membrane from the ring that was lifted away from the dissection chamber in step 11 above with a scalpel and put it in a 1.5-mL centrifuge tube with lysis buffer (i.e., 40 μL). Make sure to push the membrane at the bottom of the tube and that it is immersed in lysis buffer (see Note 15). 3.6 Protein Extraction

The optimization of the protein extraction protocol was performed to maximize the de-cross-linking of the fixative agents while keeping the lysate as concentrated as possible. In addition to the experimental samples, the following controls should also be considered: (1) a positive whole cell lysate control to compare with the proteome of the protrusion sample and (2) a negative control to determine the background proteins that are released to the media and can be found on the membrane, such as exosomes. In this case, the same number of cuts that were isolated for cellular protrusions should be isolated in empty areas surrounding cells. Finally, when setting up conditions, silver staining (e.g., ProteoSilver™ Silver Stain Kit) might be used to determine the quality of the proteins present in the isolated samples even in those with low amounts of proteins, such as the cellular protrusion samples (see Note 16). 1. Take the isolated LCM samples from steps 12 and 13 in Subheading 3.5 and thaw them if they were stored at 80  C (see Note 17). 2. Sonicate for 5 min and do a quick spin to bring all samples down.

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3. Leave 20 min on ice. Vortex and do a quick spin to bring all samples down. 4. Heat at 37  C for 30 min (see Note 18). Vortex and do a quick spin to bring all samples down. 5. Heat at 100  C for 20 min. Vortex and do a quick spin to bring all samples down. 6. Leave at 60  C for 2 h. Vortex and do a quick spin to bring all samples down. 7. Sonicate for 5 min and do a quick spin to bring all samples down. 8. The lysates are now ready for (1) RCDC protein assay if the protein concentration needs to be measured (see Note 19); (2) add Laemmli buffer to run samples in a Bis-Tris gel if a set number of cuts were obtained (see Note 20); or (3) directly proceed to step 1 of Subheading 3.8. 3.7 Sample Preparation for Mass Spectrometry Using Limited Gel

1. Use a 7.5% Mini-Protean TGX gel or, alternatively, you can polymerize an 8% acrylamide Bis-Tris gel (see Note 21). 2. Calculate the protein concentration for each sample (for example, for the orbitrap LC-MS/MS, we ran 2 and 5 μg of protein) or load a predetermined number of cuts (see Note 20). 3. Add the corresponding volume of 5 Laemmli buffer containing 150 mM DTT, vortex, and spin the samples. 4. Boil (100  C) the samples for 5 min. 5. When the samples are cool, vortex and spin again. 6. Load the samples on the acrylamide gel. 7. Run the gel at 30 mA, or about 5–10 min, down to ~1 cm inside the gel (enough to visualize the MW marker ladder separation) (see Note 22). 8. Stop running the gel and fix it for 60 min in the fixing solution (water:methanol:acetic acid ¼ 40:40:8). 9. Inside a biosafety cabinet, excise each lane carefully from the gel (see Note 23). 10. Put the sliced gel piece in a 1.5-mL centrifuge tube filled with 1% acetic acid in molecular biology grade water. The samples can be stored in this solution at 4  C until analyzed by MS. 11. Standard MS sample preparation should be performed (see Note 24). 12. Briefly, for in-gel digestion, dice the samples into 1 mm  1 mm squares, rinse multiple times with 50 mM ammonium bicarbonate, and reduce with 5 mM DTT and 50 mM ammonium bicarbonate at 55  C for 30 min.

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13. Remove the residual solvent and perform alkylation using 10 mM propionamide in 50 mM ammonium bicarbonate for 30 min at room temperature. 14. Rinse the gel pieces with 50% acetonitrile and 50 mM ammonium bicarbonate and place in a speed vacuum for 5 min. 15. Digest with trypsin/LysC overnight at 37  C. 16. Peptide extraction: Spin down the tubes and collect the solvent, which contains the peptides. Add 60% acetonitrile, 39.9% water, and 0.1% formic acid and incubate for 10–15 min. The peptide pools can be dried in a speed vacuum. 17. Reconstitute the digested peptide pools and inject onto a 100 μm I.D. C18 reversed-phase analytical column (2.4 μM Reprosil-Pur), 25–50 cm in length. The UPLC was a Waters M class, operated at 300 nL/min using a linear gradient from 4% mobile phase B to 35% B. Mobile phase A consisted of 0.2% formic acid, 5% DMSO, and water; mobile phase B was 0.2% formic acid, 5% DMSO, and acetonitrile. 18. For data collected using the Orbitrap Fusion mass spectrometer, it was set to acquire data in a data-dependent fashion selecting and fragmenting by collision-induced dissociation the most intense precursor ions optimized to maximize duty cycle. An exclusion window of 60 s was used to improve proteomic depth, and multiple charge states of the same ion were not sampled. 19. For data collected using the timsTOF Pro mass spectrometer in PASEF mode, the mobility range used was 0.7 1/K0 to 1.5 1/K0 and the ramp time was 100 ms. The effective duty cycle was 120 Hz, where the dynamic exclusion setting was set to 0.4 min. 3.8 Sample Preparation for Mass Spectrometry Using iST Sample Preparation Kit from PreOmics

As an alternative to using limited gels, the samples can be directly prepared using the iST Sample Preparation Kit from PreOmics. This allows for a fast and complete sample preparation that minimizes possible background contamination. The kit can be used with samples as small as 1 μg of protein. In our case, we were able to use it for small cellular protrusions, like filopodia. The kit includes lysis, digest, and peptide purification protocols (see Note 25). 1. Follow the manufacturer’s instructions. 2. The peptide pool is obtained at the end, ready to be run for MS.

3.9

Data Analysis

1. MS data analysis will vary depending on the mass spectrometer and software used. In our case, the MS data were analyzed using Preview and Byonic v2.6.49 as well as custom tools for

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data analysis developed in MatLab at Stanford University. Peak selection was handled automatically within Byonic. 2. MS/MS data were searched against a UniProtKB FASTA database containing 16,972 reviewed Mus musculus entries (various dates). Propionamidation (+71.037114 @ C) was set as a fixed modification, Deamination (+0.984016 @ N) and Acetylation (+42.010565 @ K) were set as common1 modifications, Oxidation (+15.994915 @ M) was set as common2 modification, and Acetylation (+42.010565 @ Protein N-term) and Methylation (+14.01565 @ K, R) were set as rare1 modifications. Byonic was set to allow a maximum of two common modifications and one rare modification and to allow a maximum of two missed cleavages. MS/MS spectra were matched with a tolerance of 12 ppm on precursor mass and 0.4 Da on fragment mass. 3. Common contaminants were filtered automatically by Byonic and include TRYP_PIG, ALBU_BOVIN, ALBU_HUMAN, CASB_BOVIN, CASK_BOVIN, CAS1_BOVIN, CO3_HUMAN, HBA_HUMAN, HBB_HUMAN, K1M1_SHEEP, K2C1_HUMAN, K22E_HUMAN, K1C10_HUMAN, K1C15_SHEEP, K1C9_HUMAN, KRHB1_HUMAN, KRHB3_HUMAN, KRHB5_HUMAN, KRHB6_HUMAN, and TRFE_HUMAN. 4. Using Byonic, the proteome was searched with a reverse-decoy strategy and all data were filtered and presented at a 1% false discovery rate. Byonic calculates a Byonic score that is an indicator of the correctness of our peptide-spectrum matches (PSM). Byonic scores reflect the absolute quality of the PSM. Byonic scores range from 0 to 1000, with 300 being a good score, 400 a very good score, and scores over 500 reflecting near-perfect matches. For our data, all filtered protein identification hits have an FDR rate lower or equal to 2.5%, a Byonic score greater than 250, and a log probability greater than 3. 5. To further discriminate nonsignificant to statistically significant proteins, we also recommend running a statistical power analysis [32]. 6. Once the statistically significant proteins are determined, a spectral counting method known as normalized spectral abundance factor (NSAF) [33] can be used to determine the relative abundance of proteins in the samples [27, 33, 34], and to verify the reproducibility, between replicates, of the quantitative data through a correlation analysis [34]. 7. Once the statistically significant proteins are determined, along with their relative abundance, a number of analyses can be performed to look at functional and/or localization differences

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[27]. The type of analysis will depend on the experimental setup and research questions and should be determined by each investigator. 8. A number of programs can be used to do these analyses including R, JMP, SSPS, or Matlab for statistical analysis; the R package SuperExactTest for the statistical analysis and visualization of multiset intersections [35]; DAVID [36, 37], Proteomaps [38], Panther [39], or GOrilla [40, 41] for gene ontology enrichment analysis; and COMPLEAT for a subcellular localization enrichment analysis [27].

4

Notes 1. All fixation solutions must be made fresh, just prior to using them. Do not store the solutions. 2. DTBP needs to be stored at 4  C but equilibrated to room temperature prior to use (30 min). The stock solution of 50 mM will turn cloudy in approximately 5 min. Use the stock before cloudiness occurs; otherwise, discard it. Reaction pH is critical; basic conditions (pH 8–10) favor mechanism of action of cross-linking by DTBP. It must be made fresh. Do not store the solution. 3. RIPA buffer (without DTT and protease inhibitors) can be stored at 4  C. However, DTT and protease inhibitors should be added freshly before use. 4. 5 Laemmli buffer can be stored for several months at 20  C, but DTT should be added freshly before use. 5. Can be stored at room temperature for extended time but protected from light. It might start to turn yellow over time, but this color change does not affect its use. 6. Cells must adhere well to the membrane in order to isolate them and/or cellular protrusions by LCM. However, whether you use fibronectin, poly-lysine, collagen, or any other extracellular matrix protein coatings, you need to make sure that it does not hinder the formation of the cellular protrusion of interest. 7. MMI suggests that you seed the cells in the ring and later move it to the UV-permeable microdissection chamber. However, the most important issue when using an MMI live chamber is to ensure that there is no leakage once the ring is attached to the microdissection chamber. If the ring leaks and wets the bottom of the membrane prior to putting inside the microdissection chamber, it will not properly attach to the adhesive area of the chamber. As a result, the LCM-isolated cut will not stick to it and will not be collected after removal of the ring. For this

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reason, we always attach the ring to the microdissection chamber first and only after asserting that it does not leak, we seed the cells. It is important to minimize bubbles and wrinkles when setting the ring in the microdissection chamber. 8. Do not disturb the cells by moving the dish after seeding. Leave the cells in the incubator so that they can attach to the membrane and make protrusions. The time of adhesion and formation of cellular protrusions depends on both the cell types used and the types of cellular protrusions of interest. In our case, we incubate CAD cells for 2–3 h, when looking for filopodia and/or TNTs but waited up to 10 days with media change every 2–3 days for axons and dendrites in order to allow for cell differentiation upon serum starvation. The time must be predetermined by each investigator based on their cell/protrusion types. 9. Many different ways to induce cell protrusions have been published. Test different substances or physical conditions (i.e., hypoxia) that can work with your cell type to increase the number of cellular protrusions you are interested in. Remember that for each treatment, whole cell control samples should be acquired. 10. The proper stimuli and/or concentrations should be predetermined by the investigator based on the cell type used and cellular protrusions of interest. For instance, prolong H2O2 exposure and/or high concentration can be harmful to the cells and could result in the cells detaching from the membrane. In addition, to avoid disturbing the cells, care should be taken when adding and aspirating solutions. 11. Time of cross-linking is critical. Proceed to the next step immediately to avoid over-cross-linking. 12. All the procedures with the LCM must be done with filtered PBS in the dish to avoid protein degradation [26]. 13. Because time degrades proteins, longer periods of cutting must be avoided (see Fig. 1d). 14. If your cuts are limited to a specific area, it helps if you outline with a marker on the bottom of the dissection chamber the desired area. 15. Both protrusion and membrane samples can be stored at 80  C for several weeks. 16. Silver stain is not compatible with some types of MS. 17. Since the samples are in small volumes, the caps of the centrifuge tubes should be tightly closed and surrounded with parafilm to avoid evaporation and drying of the samples.

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18. DTBP-cross-linked proteins can be cleaved by reducing the disulfide bond of the spacer arm with 100–150 mM DTT at 37  C for 30 min (see reagent setup), thus freeing the isolated proteins from the cross-linked product. 19. RCDC is compatible with detergents (such as SDS) and reducing agents (such as DTT). Bradford and BCA are compatible with reducing agents and detergents, respectively. Make a standard curve with BSA and use as blank the protein extraction supernatant of a membrane from a prepared MMI live chamber dish but never seeded with cells. Fibronectin is a protein, and for that reason, it is likely that fibronectin coating increases protein results. 20. RCDC can be less reliable for very small samples of isolated cellular protrusions, and it takes away a large amount of the isolated samples. In order to use all of the LCM-isolated cellular protrusions for MS analysis, we recommend determining what the total number of cuts required is and avoid using the RCDC. The number of cuts required will vary depending on the cellular protrusion types and should be predetermined by each investigator. When using the orbitrap LC-MS/MS, we ran 2 and 5 μg of total protein or 3,000 to 6,000 cuts, depending on the types of cellular protrusions isolated. When using the timsTOF Pro mass spectrometer, the number of cuts was reduced due to the greater sensitivity of this mass spectrometer compared to the Orbitrap. 21. To avoid keratin contamination, all these steps and the required solutions should be done in a biological hood and with molecular grade water. 22. This “limited gel” allows for the separation of the proteins by size in preparation for MS, and each lane can be run as one sample by MS. 23. It is easier to cut the samples if they are loaded between stained ladders. 24. Because of the minute sample size, only very sensitive mass spectrometers should be used. We recommend using either the Thermo Scientific Orbitrap Fusion mass spectrometer or the timsTOF Pro mass spectrometer from Bruker. 25. The iST sample preparation takes about 2½ h and includes all required steps, such as lysis to denature, reduce, and alkylate proteins, digestion using a LysC and Trypsin mix, and peptide purification, which includes removal of hydrophilic contaminants.

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Acknowledgments This work was supported by a 2014 CSUPERB New Investigator Grant and the National Institute of General Medical Sciences of the National Institutes of Health under Award Number SC2GM111144 awarded to K.G. References 1. Hanson JC, Tangrea MA, Kim S et al (2011) Emmert-Buck, Expression microdissection adapted to commercial laser dissection instruments. Nat Protoc 6:457–467 2. Gru¨newald A, Rygiel KA, Hepplewhite PD et al (2016) Mitochondrial DNA depletion in respiratory chain-deficient Parkinson disease neurons. Ann Neurol 79:366–378 3. Drummond ES, Nayak S, Ueberheide B, Wisniewski T (2015) Proteomic analysis of neurons microdissected from formalin-fixed, paraffin-embedded Alzheimer’s disease brain tissue. Sci Rep 5:15456 4. Nawandar DM, Wang A, Makielski K et al (2015) Differentiation-dependent KLF4 expression promotes lytic Epstein-Barr virus infection in epithelial cells. PLoS Pathog 11: e1005195 5. Pflugradt R, Schmidt U, Landenberger B et al (2011) A novel and effective separation method for single mitochondria analysis. Mitochondrion 11:308–314 6. Zivraj KH, Tung YC, Piper M (2010) Subcellular profiling reveals distinct and developmentally regulated repertoire of growth cone mRNAs. J Neurosci 30:15464–15478 7. Ezzoukhry Z, Henriet E, Cordelie`res FP (2018) Combining laser capture microdissection and proteomics reveals an active translation machinery controlling invadosome formation. Nat Commun 9:2031 8. Gutstein HB, Morris JS, Annangudi SP, Sweedler JV (2008) Microproteomics: analysis of protein diversity in small samples. Mass Spectrom Rev 27:316–330 9. Gambade A, Zreika S, Gue´guinou M et al (2016) Activation of TRPV2 and BKCa channels by the LL-37 enantiomers stimulates calcium entry and migration of cancer cells. Oncotarget. https://doi.org/10.18632/ oncotarget.8122 10. Costanzo M, Abounit S, Marzo L et al (2013) Transfer of polyglutamine aggregates in neuronal cells occurs in tunneling nanotubes. J Cell Sci 126:3678–3685

11. Mo¨ller J, Lu¨hmann T, Chabria M et al (2013) Macrophages lift off surface-bound bacteria using a filopodium-lamellipodium hook-andshovel mechanism. Sci Rep 3:2884 12. Dent EW, Gupton SL, Gertler FB (2011) The growth cone cytoskeleton in axon outgrowth and guidance. Cold Spring Harb Perspect Biol 3:a001800 13. Van Audenhove I, Denert M, Boucherie C et al (2016) Fascin rigidity and L-plastin flexibility cooperate in cancer cell invadopodia and filopodia. J Biol Chem 291:9148–9160 14. Gousset K, Schiff E, Langevin C et al (2009) Prions hijack tunnelling nanotubes for intercellular spread. Nat Cell Biol 11:328–336 15. Eugenin EA, Gaskill PJ, Berman JW (2009) Tunneling nanotubes (TNT) are induced by HIV-infection of macrophages: a potential mechanism for intercellular HIV trafficking. Cell Immunol 254:142–148 16. Thayanithy V, Dickson EL, Steer C et al (2014) Tumor-stromal cross talk: direct cell-to-cell transfer of oncogenic microRNAs via tunneling nanotubes. Transl Res 164:359–365 17. Brayford S, Bryce NS, Schevzov G et al (2016) Tropomyosin promotes lamellipodial persistence by collaborating with Arp2/3 at the leading edge. Curr Biol 26(10):1312–1318 18. Sherer NM, Mothes W (2008) Cytonemes and tunneling nanotubules in cell-cell communication and viral pathogenesis. Trends Cell Biol 18:414–420 19. Thomsen R, Lade Nielsen A (2011) A Boyden chamber-based method for characterization of astrocyte protrusion localized RNA and protein. Glia 59:1782–1792 20. Kadiu I, Gendelman HE (2011) Human immunodeficiency virus type 1 endocytic trafficking through macrophage bridging conduits facilitates spread of infection. J Neuroimmune Pharmacol 6:658–675 21. Mili S, Moissoglu K, Macara IG (2008) Genome-wide screen identifies localized RNAs anchored at cell protrusions through microtubules and APC. Nature 453:115–119

Proteome of Cellular Protrusions Identified by LCM/MS 22. Mimae T, Ito A (2015) New challenges in pseudopodial proteomics by a laser-assisted cell etching technique. Biochim Biophys Acta 1854:538–546 23. Gousset K, Marzo L, Commere P, Zurzolo C (2013) Myo10 is a key regulator of TNT formation in neuronal cells. J Cell Sci 126:4424–4435 24. Eltoum I, Fredenburgh J, Myers RB, Grizzle WE (2001) Introduction to the theory and practice of fixation of tissues. J Histotechnol 24:173–190 25. Gosselin MA, Guo W, Lee RJ (2001) Efficient gene transfer using reversibly cross-linked low molecular weight polyethylenimine. Bioconjug Chem 12:989–994 26. Gordon A, Kannan SK, Gousset K (2018) A novel cell fixation method that greatly enhances protein identification in microproteomic studies using laser capture microdissection and mass spectrometry. Proteomics 18: e1700294 27. Gousset K, Gordon A, Kumar Kannan S, Tovar J (2019) A novel microproteomic approach using laser capture microdissection to study cellular protrusions. Int J Mol Sci 20:1172 28. Osswald M, Jung E, Sahm F et al (2015) Brain tumour cells interconnect to a functional and resistant network. Nature 528:93–98 29. Desir S, Dickson EL, Vogel RI et al (2016) Tunneling nanotube formation is stimulated by hypoxia in ovarian cancer cells. Oncotarget. https://doi.org/10.18632/oncotarget.9504 30. Hashimoto M, Bhuyan F, Hiyoshi M et al (2016) Potential role of the formation of tunneling nanotubes in HIV-1 spread in macrophages. J Immunol 196:1832–1841 31. Victoria GS, Arkhipenko A, Zhu S et al (2016) Astrocyte-to-neuron intercellular prion transfer is mediated by cell-cell contact. Sci Rep 6:20762

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32. Levin Y (2011) The role of statistical power analysis in quantitative proteomics. Proteomics 11:2565–2567. https://doi.org/10.1002/ pmic.201100033 33. Arike L, Peil L (2014) Spectral counting labelfree proteomics. Methods Mol Biol 1156:213–222 34. McIlwain S, Mathews M, Bereman MS, Rubel EW, MacCoss MJ, Noble WS (2012) Estimating relative abundances of proteins from shotgun proteomics data. BMC Bioinformatics 13:308. https://doi.org/10.1186/14712105-13-308 35. Wang M, Zhao Y, Zhang B (2015) Efficient test and visualization of multiset intersections. Sci Rep 5:16923 36. Huang DW, Sherman BT, Lempicki RA (2009) Systematic and integrative analysis of large gene lists using DAVID Bioinformatics Resources. Nat Protoc 4:44–57 37. Huang DW, Sherman BT, Lempicki RA (2009) Bioinformatics enrichment tools: paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res 37:1–13 38. Liebermeister W, Noor E, Flamholz A, Davidi D, Bernhardt J, Milo R (2014) Visual account of protein investment in cellular functions. Proc Natl Acad Sci U S A 111:8488–8493 39. Mi H, Muruganujan A, Ebert D, Huang X, Thomas PD (2019) PANTHER version 14: more genomes, a new PANTHER GO-slim and improvements in enrichment analysis tools. Nucleic Acids Res 47:D419–D426. https://doi.org/10.1093/nar/gky1038 40. Eden E, Navon R, Steinfeld I, Lipson D, Yakhini Z (2009) GOrilla: a tool for discovery and visualization of enriched GO terms in ranked gene lists. BMC Bioinformatics 10:48 41. Eden E, Lipson D, Yogev S, Yakhini Z (2007) Discovering motifs in ranked lists of DNA sequences. PLoS Comput Biol 3:e39

Part II Shotgun Proteomics of Non-model Organisms

Chapter 4 Isolation of Apoplastic Fluid from Woody Plant Leaves: Grapevine and Coffee as a Case Study Andreia Figueiredo and Leonor Guerra-Guimara˜es Abstract Proteomics is one of the key approaches to understand plant cell physiology involving the regulation of expression of many genes and metabolite production. Technical advances allowed a deeper characterization of plant proteomes, highlighting the need to study cellular compartments. The apoplast is the cellular compartment external to the plasma membrane including the cell wall, where a broad range of processes take place including intercellular signaling, metabolite transport, and plant–microbe interactions. Due to the fragile nature of leaf tissues, it is a challenge to obtain apoplastic fluids from leaves while maintaining cell integrity, which is particularly true for woody plants. Here, we describe the vacuum infiltrationcentrifugation (VIC) method for the extraction of the apoplastic fluid compatible with high-throughput proteomic approaches and biochemical analysis from different woody plants. Key words VIC method, Apoplast, Apoplastic fluid, Grapevine leaves, Coffee leaves, Vitis vinifera, Coffea arabica, Electrophoresis, Concentration

1

Introduction The plant extracellular space, or apoplast, is a highly dynamic compartment affected by environmental conditions and participates in plant signaling, defense, growth, physiology, cell wall maintenance, and reproduction [1]. It is defined by the plant cell wall, extracellular matrix, and the intercellular spaces where the apoplastic fluid (APF) circulates [2]. APF studies allow uncovering plant’s secretome. It comprehends a set of proteins and small molecules exported out of the symplast that play several roles, where the response to biotic and abiotic stresses takes a spotlight position [3–5]. Apoplastic fluid extraction from plant systems is far from easy and remains challenging. This is particularly true for recalcitrant plants, as the majority of them are woody plants. Thus, the study of apoplast dynamics is hindered by low protein abundance and high possibility of damaging the plant cell, leading to cytoplasmic

Mo´nica Carrera and Jesu´s Mateos (eds.), Shotgun Proteomics: Methods and Protocols, Methods in Molecular Biology, vol. 2259, https://doi.org/10.1007/978-1-0716-1178-4_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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content contamination. The selection of an appropriate extraction protocol is a crucial step in a proteomic approach, and subsequent compatibility with the analytical techniques (e.g., mass spectrometry) has to be considered. The most commonly used technique for plant APF extraction is the vacuum infiltration-centrifugation (VIC), described by Klement in 1965 [6]. The VIC involves two critical steps: vacuum infiltration with appropriate extraction buffer and centrifugation [7]. The composition of the infiltration buffer is one of the key features of this method, as it should combine the maintenance of osmotic pressure to prevent plasma membrane damage and cytoplasm content leaking, and the efficiency to extract cell wall proteins [7]. High concentrations of ionic reagents, such as KCl, NaCl, or LiCl, are normally considered to guarantee weakly bound cell wall protein solubilization [2]. The centrifugation step is also limiting, as centrifugation speed may cause damage to cell walls and membranes [7]. Here, we present a vacuum protocol optimized for woody plant leaves, namely Vitis vinifera and Coffea arabica. This protocol is relatively easy to perform, allows obtaining an acceptable protein yield, avoids contamination with cytosolic components, and is compatible with high-throughput mass spectrometry-based proteomic approaches, namely shotgun proteomics. We discuss critical points within the extraction procedure, namely the influence of the composition of the infiltration buffer. Using this protocol, we have isolated apoplastic proteins from both coffee and grapevine leaves [8–10], yielding a good coverage of the apoplast proteome.

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ cm at 25  C) and analytical-grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations.

2.1

Plant Material

1. Field or greenhouse-grown plants of appropriate age and life stage, preferably with fully expanded leaves, should be used (see Notes 1 and 2). The protocol is optimized for grapevine (Vitis vinifera) and coffee (Coffea arabica) leaves. Approximately 25 g of fresh weight was used for grapevine (around 30 fully expanded leaves), and about 10 g of fresh weight was used for coffee (around 8 pairs of leaves). 2. Paper towel. 3. Scalpel.

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4. Glass beaker. 5. Parafilm. 6. Weighing balance. 2.2 Apoplastic Fluid Isolation Buffers and Equipment

1. Ice-cold deionized water: Store several 1-L glass bottles at 4  C. 2. Coffee infiltration buffer: 100 mM Tris–HCl buffer, pH 7.6; add about 100 mL of water to a 1-L graduated a glass beaker, weigh 121.14 g of Tris and transfer to the beaker, add water to a volume of 900 mL, and mix with a magnetic stir bar till it dissolves. Add 500 mM L-ascorbic acid (88.06 g) and 500 mM potassium chloride (37.28 g) and mix till it dissolves (see Note 3). Adjust pH with HCl. Make up to 1 L with water. Keep it at 4  C for no longer than 2 weeks. Add 25 mM 2-mercaptoethanol (1.7 mL per 100 mL of buffer) to the buffer solution just before use (see Note 4) [8, 9]. 3. Grapevine infiltration buffer: 100 mM Tris–HCl buffer, pH 7.6; add about 100 mL of water to a 1-L graduated a glass beaker, weigh 121.14 g of Tris and transfer to the beaker, add water to a volume of 900 mL, and mix with a magnetic stir bar till it dissolves. Adjust pH with HCl. Add 500 mM potassium chloride (37.275 g) and 6 mM CHAPS (3.68 g) (see Note 5) and mix till it dissolves. Make up to 1 L with water. Keep it at 4  C. Add 2% of sodium sulfite (20 g/L) to the buffer solution just before use (see Note 6) [10, 11]. 4. Soft towel paper. 5. Large glass Kitasato flask, or vacuum flask, with lid. 6. Vacuum pump to apply vacuum at 25 kPa (187.52 mmHg). 7. 20-mL plastic syringe without a plunger. 8. 50-mL falcon tube or equivalent. 9. 1.5-mL Eppendorf tubes. 10. Refrigerated centrifuge with fixed angle rotor for 50-mL centrifuge tubes.

2.3 Apoplast Protein Concentration and Quantification

1. Refrigerated centrifuge with fixed angle rotor for 15 mL tubes. 2. Centrifugal filter Vivaspin2. 3. Eppendorf tubes (1.5 mL). 4. Micropipettes. 5. Bradford reagent. 6. Protein standard: 2.5 mg/mL albumin in water. 7. 96 ELISA well microplates. 8. ELISA reader (ABS 595 nm).

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3 3.1

Andreia Figueiredo and Leonor Guerra-Guimara˜es

Methods Infiltration

1. Harvest fully expanded leaves from grapevine and coffee shoots to obtain approximately 25 g and 10 g of fresh weight material, respectively. Remove the middle and lateral veins using a scalpel (see Fig. 1a, b, Note 7). 2. Cut leaves into pieces of around 2 cm2 to a glass or plastic beaker (see Fig. 1, Note 8) kept on ice. Transfer the leaf pieces to a large glass Kitasato flask filled with ice-cold infiltration buffer (see Fig. 2a, Note 9). 3. Apply vacuum to the Kitasato flask employing a vacuum pump. A final pressure of about 25 kPa around 187.52 mmHg is sufficient for infiltration. A total infiltration time of 3 min (six periods of 30 s) was applied. 4. Slowly release the vacuum by releasing the vacuum tube on the Kitasato flask. During vacuum release, the leaves are slowly infiltrated with the infiltration buffer. During this process, leaf tissue becomes water-soaked and dark in color (see Fig. 2b).

Fig. 1 Preparation of grapevine (a) and coffee (b) leaves for infiltration. Leaves are cut with a scalpel in 2 cm2 squares after removing the middle and lateral veins (grapevine, c) and the middle vein (coffee, d)

Extraction of Apoplastic Fluid from Woody Plants

53

Fig. 2 Vacuum is applied to the Kitasato flask (a). Leaf squares are infiltrated, becoming water-soaked and presenting dark punctuations (b) 3.2 Isolation of Apoplastic Fluid by Centrifugation

1. After infiltration, rinse leaf pieces twice in ice-cold deionized water (see Fig. 3a, b). Dry the leaf pieces in soft towel paper (see Fig. 3c). 2. Stack up the infiltrated leaves into a 20-mL syringe and then in a 50-mL centrifuge tube. Inside the centrifuge tube, place a 1.5-mL Eppendorf, and the tip of the syringe is oriented to be inside the Eppendorf to collect the APF (see Fig. 3e). 3. Place the tubes inside the centrifuge and spin at 5000  g for 15 min at 4  C (see Fig. 3f). 4. During centrifugation, the APF is extruded from the leaves and collected in the Eppendorf tube located in the bottom of the centrifuge tube (see Fig. 3g). Immediately after recovery, the APF should be stored at 20  C until further use.

3.3 Apoplast Protein Concentration and Quantification

1. Defreeze the APF samples and centrifuge at 10,000  g for 10 min at 4  C. After supernatant recovery, desalt and concentrate the APF on centrifugal filter Vivaspin2 according to the manufacturer’s instructions (Sartorius). This ultrafiltration process uses anisotropic semipermeable membranes to separate macromolecular species and solvents by size. If a nondenaturing buffer/solution (e.g., glycerol 10%) is used for sample desalt, APF extracts can be considered for enzymatic activity assays, namely malate dehydrogenase activity (to assess cytoplasmic contamination, see Note 10). 2. Determine APF protein dye-binding method [12] procedure. A microassay low-concentration protein

quantification by the Bradford using the Bio-Rad protein assay 96-well microplate protocol for samples (