2D PAGE. Sample Preparation and Fractionation

2D PAGE: Sample Preparation and Fractionation M E T H O D S I N M O L E C U L A R B I O L O G YTM John M. Walker,...

Author: Anton Posch

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2D PAGE: Sample Preparation and Fractionation

M E T H O D S

I N

M O L E C U L A R

B I O L O G YTM

John M. Walker, SERIES EDITOR 460. Essential Concepts in Toxicogenomics, edited by Donna L. Mendrick and William B. Mattes, 2008 459. Prion Protein Protocols, edited by Andrew F. Hill, 2008 458. Artificial Neural Networks: Methods and Applications, edited by David S. Livingstone, 2008 457. Membrane Trafficking, edited by Ales Vancura, 2008 456. Adipose Tissue Protocols, Second Edition, edited by Kaiping Yang, 2008 455. Osteoporosis, edited by Jennifer J. Westendorf, 2008 454. SARS- and Other Coronaviruses: Laboratory Protocols, edited by Dave Cavanagh, 2008 453. Bioinformatics, Volume 2: Structure, Function, and Applications, edited by Jonathan M. Keith, 2008 452. Bioinformatics, Volume 1: Data, Sequence Analysis, and Evolution, edited by Jonathan M. Keith, 2008 451. Plant Virology Protocols: From Viral Sequence to Protein Function, edited by Gary Foster, Elisabeth Johansen, Yiguo Hong, and Peter Nagy, 2008 450. Germline Stem Cells, edited by Steven X. Hou and Shree Ram Singh, 2008 449. Mesenchymal Stem Cells: Methods and Protocols, edited by Darwin J. Prockop, Douglas G. Phinney, and Bruce A. Brunnell, 2008 448. Pharmacogenomics in Drug Discovery and Development, edited by Qing Yan, 2008 447. Alcohol: Methods and Protocols, edited by Laura E. Nagy, 2008 446. Post-translational Modification of Proteins: Tools for Functional Proteomics, Second Edition, edited by Christoph Kannicht, 2008 445. Autophagosome and Phagosome, edited by Vojo Deretic, 2008 444. Prenatal Diagnosis, edited by Sinhue Hahn and Laird G. Jackson, 2008 443. Molecular Modeling of Proteins, edited by Andreas Kukol, 2008 442. RNAi: Design and Application, edited by Sailen Barik, 2008 441. Tissue Proteomics: Pathways, Biomarkers, and Drug Discovery, edited by Brian Liu, 2008 440. Exocytosis and Endocytosis, edited by Andrei I. Ivanov, 2008 439. Genomics Protocols, Second Edition, edited by Mike Starkey and Ramnanth Elaswarapu, 2008 438. Neural Stem Cells: Methods and Protocols, Second Edition, edited by Leslie P. Weiner, 2008 437. Drug Delivery Systems, edited by Kewal K. Jain, 2008 436. Avian Influenza Virus, edited by Erica Spackman, 2008 435. Chromosomal Mutagenesis, edited by Greg Davis and Kevin J. Kayser, 2008

434. Gene Therapy Protocols: Volume 2, Design and Characterization of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 433. Gene Therapy Protocols: Volume 1, Production and In Vivo Applications of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 432. Organelle Proteomics, edited by Delphine Pflieger and Jean Rossier, 2008 431. Bacterial Pathogenesis: Methods and Protocols, edited by Frank DeLeo and Michael Otto, 2008 430. Hematopoietic Stem Cell Protocols, edited by Kevin D. Bunting, 2008 429. Molecular Beacons: Signalling Nucleic Acid Probes, Methods and Protocols, edited by Andreas Marx and Oliver Seitz, 2008 428. Clinical Proteomics: Methods and Protocols, edited by Antonio Vlahou, 2008 427. Plant Embryogenesis, edited by Maria Fernanda Suarez and Peter Bozhkov, 2008 426. Structural Proteomics: High-Throughput Methods, edited by Bostjan Kobe, Mitchell Guss, and Huber Thomas, 2008 425. 2D PAGE: Sample Preparation and Fractionation, Volume 2, edited by Anton Posch, 2008 424. 2D PAGE: Sample Preparation and Fractionation, Volume 1, edited by Anton Posch, 2008 423. Electroporation Protocols, edited by Shulin Li, 2008 422. Phylogenomics, edited by William J. Murphy, 2008 421. Affinity Chromatography, Methods and Protocols, Second Edition, edited by Michael Zachariou, 2007 420. Drosophila, Methods and Protocols, edited by Christian Dahmann, 2008 419. Post-Transcriptional Gene Regulation, edited by Jeffrey Wilusz, 2008 418. Avidin-Biotin Interactions, Methods and Applications, edited by Robert J. McMahon, 2008 417. Tissue Engineering, Second Edition, edited by Hannsjörg Hauser and Martin Fussenegger, 2007 416. Gene Essentiality: Protocols and Bioinformatics, edited by Andrei L. Osterman, 2008 415. Innate Immunity, edited by Jonathan Ewbank and Eric Vivier, 2007 414. Apoptosis and Cancer: Methods and Protocols, edited by Gil Mor and Ayesha B. Alvero, 2008 413. Protein Structure Prediction, Second Edition, edited by Mohammed Zaki and Chris Bystroff, 2008 412. Neutrophil Methods and Protocols, edited by Mark T. Quinn, Frank R. DeLeo, and Gary M. Bokoch, 2007 411. Reporter Genes for Mammalian Systems, edited by Don Anson, 2007 410. Environmental Genomics, edited by Cristofre C. Martin, 2007 409. Immunoinformatics: Predicting Immunogenicity In Silico, edited by Darren R. Flower, 2007

M E T H O D S I N M O L E C U L A R B I O L O G YT M

2D PAGE: Sample Preparation and Fractionation Volume 2

Edited by

Anton Posch Bio-Rad Laboratories GmbH, Munich, Germany

Editor Anton Posch Bio-Rad Laboratories GmbH Munich, Germany Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Herts., UK

ISBN: 978-1-60327-209-4 ISSN: 1064-3745

e-ISBN: 978-1-60327-210-0

Library of Congress Control Number: 2007943017 ©2008 Humana Press, a part of Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: Figure 3, Chapter 14, “The Terminator: A Device for High Throughput Extraction of Plant Material,” by B. M. van den Berg. Printed on acid-free paper 987654321 springer.com

Preface

This book, split into two volumes, presents a broad coverage of the principles and recent developments of sample preparation and fractionation tools in Expression Proteomics in general and for two-dimensional electrophoresis (2-DE) in particular. 2-DE, with its unique capacity to resolve thousands of proteins in a single run, is still a fundamental research tool for nearly all protein-related scientific projects. The methods described here in detail are not limited to 2-DE and can also be applied to other protein separation techniques. Because each biological sample is unique, a suited sample preparation strategy has to consider the type of sample as well as the type of biological question being addressed. The complex nature of proteins often requires a multitude of sample preparation options. In addition, sample preparation is not only a prerequisite for a successful and reproducible Proteomics experiment, but also the key factor to meaningful data evaluation. Interestingly, not much attention was paid to this area during Proteomics methodology development and therefore this book is intended to explain in depth how proteins from various sources can be properly isolated and prepared for reproducible Proteome analysis. The application of fractionation and enrichment strategies has become a major part of sample preparation. The number of possible different proteins in a cell or tissue sample is believed to be in the several hundreds of thousands, spanning concentration ranges from the level of a single molecule to micromolar amounts, and no single analytical method developed today is capable of resolving and detecting such a diverse sample. Sample fractionation reduces the overall complexity of the sample, and enriches low abundance proteins relative to the original sample. Proteins that may originally have been undetectable are thus rendered amenable to analysis by 2-DE and a broad variety of gel-free mass spectrometry-based technologies. This book is for students of Biochemistry, Biomedicine, Biology, and Genomics and will be an invaluable source for the experienced, practicing scientist, too. Anton Posch v

Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

v xi

1.

Application of Fluorescence Dye Saturation Labeling for Differential Proteome Analysis of 1,000 Microdissected Cells from Pancreatic Ductal Adenocarcinoma Precursor Lesions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Barbara Sitek, Bence Sipos, Günter Klöppel, Wolff Schmiegel, Stephan A. Hahn, Helmut E. Meyer, and Kai Stühler 2. Albumin and Immunoglobulin Depletion of Human Plasma . . . . . . . . 15 Rosalind E. Jenkins, Neil R. Kitteringham, Carrie Greenough, and B. Kevin Park 3. Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase Chromatography of Human Serum . . . . . . . . . . 27 James Martosella and Nina Zolotarjova 4. Immunoafﬁnity Fractionation of Plasma Proteins by Chicken IgY Antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 Lei Huang and Xiangming Fang 5.

Proteomics of Cerebrospinal Fluid: Methods for Sample Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 John E. Hale, Valentina Gelfanova, Jin-Sam You, Michael D. Knierman, and Robert A. Dean 6. Sample Preparation of Bronchoalveolar Lavage Fluid . . . . . . . . . . . . . . . 67 Baptiste Leroy, Paul Falmagne, and Ruddy Wattiez 7. Preparation of Nasal Secretions for Proteome Analysis . . . . . . . . . . . . . . 77 Begona Casado, Paolo Iadarola, and Lewis K. Pannell 8.

Preparation of Urine Samples for Proteomic Analysis . . . . . . . . . . . . . . . 89 Rembert Pieper 9. Isolation of Cytoplasmatic Proteins from Cultured Cells for Two-Dimensional Gel Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Ying Wang, Jen-Fu Chiu, and Qing-Yu He

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10.

Sample Preparation of Culture Medium from Madin-Darby Canine Kidney Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daniel Ambort, Daniel Lottaz, and Erwin Sterchi 11. Sample Preparation for Mass Spectrometry Analysis of Formalin-Fixed Parafﬁn-Embedded Tissue: Proteomic Analysis of Formalin-Fixed Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicolas A. Stewart and Timothy D. Veenstra 12. Metalloproteomics in the Molecular Study of Cell Physiology and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hermann-Josef Thierse, Stefanie Helm, and Patrick Pankert 13. Protein Extraction from Green Plant Tissue . . . . . . . . . . . . . . . . . . . . . . . . .

113

131

139

149

Ragnar Flengsrud 14. The Terminator: A Device for High-Throughput Extraction of Plant Material . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 B. M. van den Berg 15.

Isolation of Mitochondria from Plant Cell Culture. . . . . . . . . . . . . . . . . . . 163

Etienne H. Meyer and A. Harvey Millar 16. Isolation and Preparation of Chloroplasts from Arabidopsis thaliana Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 Sybille E. Kubis, Kathryn S. Lilley, and Paul Jarvis 17.

Isolation of Plant Cell Wall Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187

Elisabeth Jamet, Georges Boudart, Gisèle Borderies, Stephane Charmont, Claude Laﬁtte, Michel Rossignol, Herve Canut, and Rafael Pont-Lezica 18. Isolation and Fractionation of the Endoplasmic Reticulum from Castor Bean (Ricinus communis) Endosperm for Proteomic Analyses. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 203 William J. Simon, Daniel J. Maltman and Antoni R. Slabas 19.

Cell Wall Fractionation for Yeast and Fungal Proteomics . . . . . . . . . . . 217 Aida Pitarch, César Nombela, and Concha Gil 20. Collection of Proteins Secreted from Yeast Protoplasts in Active Cell Wall Regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 Aida Pitarch, César Nombela, and Concha Gil 21. Sample Preparation Procedure for Cellular Fungi . . . . . . . . . . . . . . . . . . . 265 Alois Harder

Contents 22.

23.

24.

25.

ix

Isolation and Enrichment of Secreted Proteins from Filamentous Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Martha L. Medina and Wilson A. Francisco Isolation and Solubilization of Cellular Membrane Proteins from Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 Kheir Zuobi-Hasona and L. Jeannine Brady Isolation and Solubilization of Gram-Positive Bacterial Cell Wall-Associated Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295 Jason N. Cole, Steven P. Djordjevic, and Mark J. Walker

Cell Fractionation of Parasitic Protozoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Wanderley de Souza, José Andrés Morgado-Diaz, and Narcisa L. Cunha-e-Silva Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333

Contributors

Daniel Ambort • University of Berne, Berne, Switzerland Leroy Baptiste • University of Mons-Hainaut, Mons, Belgium Gisèle Borderies • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, Castanet-Tolosan, France Georges Boudart • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, Castanet-Tolosan, France L. Jeannine Brady • University of Florida, Gainesville, Florida Herve Canut • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, Castanet-Tolosan, France Begona Casado • Swiss Federal Institute of Technology, Zurich, Switzerland Stephane Charmont • Novartis Pharma AG, Basel, Switzerland Jen-Fu Chiu • The University of Hong Kong, Hong Kong, China Jason N. Cole • University of Wollongong, Wollongong, Australia Narcisa L. Cunha-e-Silva • Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil Wanderley de Souza • Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil Robert A. Dean • Lilly Research Laboratories, Indianapolis, Indiana Steven P. Djordjevic • Elizabeth Macarthur Agricultural Institute, Menangle, Australia Paul Falmagne • University of Mons-Hainaut, Mons, Belgium Xiangming Fang • GenWay Biotech, Inc., San Diego, California Ragnar Flengsrud • Norwegian University of Life Sciences, Ås, Norway Wilson A. Francisco • Arizona State University, Tempe, Arizona Valentina Gelfanova • Lilly Research Laboratories, Greenfield, Indiana Concha Gil • Complutense University of Madrid, Madrid, Spain Carrie Greenough • University of Liverpool, Liverpool, Great Britain Stephan A. Hahn • Ruhr-University Bochum, Bochum, Germany John E. Hale • Lilly Research Laboratories, Greenfield, Indiana Alois Harder • Toplab GmbH, Martinsried, Germany Qing-Yu He • The University of Hong Kong, Hong Kong, China Stefanie Helm • University of Heidelberg, Mannheim, Germany Lei Huang • GenWay Biotech Inc., San Diego, California xi

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Paolo Iadarola • University of Pavia, Pavia, Italy Elisabeth Jamet • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, Castanet-Tolosan, France Paul Jarvis • University of Leicester, Leicester, Great Britain Rosalind E. Jenkins • University of Liverpool, Liverpool, Great Britain Neil R. Kitteringham • University of Liverpool, Liverpool, Great Britain Günter Klöppel • Christian Albrechts University, Kiel, Germany Michael D. Knierman • Lilly Research Laboratories, Greenfield, Indiana Sybille E. Kubis • University of Leicester, Leicester, Great Britain Claude Laﬁtte • UMR 5546 CNRS-Université Paul Sabatier-Toulouse III, Castanet-Tolosan, France Baptiste Leroy • University of Mons-Hainaut, Mons, Belgium Kathryn S. Lilley • University of Cambridge, Cambridge, Great Britain Daniel Lottaz • University of Berne, Berne, Switzerland Daniel J. Maltman • University of Durham, Durham, Great Britain James Martosella • Agilent Technologies Inc., Wilmington, Delaware Martha L. Medina • Arizona State University, Tempe, Arizona Helmut E. Meyer • Ruhr-University Bochum, Bochum, Germany Etienne H. Meyer • The University of Western Australia, Perth, Australia A. Harvey Millar • The University of Western Australia, Perth, Australia José Andrés Morgado-Diaz • Instituto Nacional de Câncer, Rio de Janeiro, Brazil César Nombela • Complutense University of Madrid, Madrid, Spain Patrick Pankert • University of Heidelberg, Mannheim, Germany Lewis K. Pannell • University of South Alabama, Mobile, Alabama B. Kevin Park • University of Liverpool, Liverpool, Great Britain Falmagne Paul • University of Mons-Hainaut, Mons, Belgium Rembert Pieper • The Institute for Genomic Research, Rockville, Maryland Aida Pitarch • Complutense University of Madrid, Madrid, Spain Rafael Pont-Lezica • UMR 5546 CNRS-Universitè Paul Sabatier-Toulouse III, Castanet-Tolosan, France Michel Rossignol • UMR 5546 CNRS-Universitè Paul Sabatier-Toulouse III, Castanet-Tolosan, France Wolff Schmiegel • Ruhr-University Bochum, Bochum, Germany William J. Simon • University of Durham, Durham, Great Britain Bence Sipos • Christian Albrechts University, Kiel, Germany Barbara Sitek • Ruhr-University Bochum, Bochum, Germany Antoni R. Slabas • University of Durham, Durham, Great Britain Erwin Sterchi • University of Berne, Berne, Switzerland Nicolas A. Stewart • National Cancer Institute at Frederick, Frederick, Maryland

Contributors

xiii

Kai Stühler • Ruhr-University Bochum, Bochum, Germany Hermann-Josef Thierse • University of Heidelberg, Mannheim, Germany B. M. van den Berg • Elexa, Enkhuizen, The Netherlands Timothy D. Veenstra • National Cancer Institute at Frederick, Frederick, Maryland Mark J. Walker • University of Wollongong, Wollongong, Australia Ying Wang • The University of Hong Kong, Hong Kong, China Ruddy Wattiez • University of Mons-Hainaut, Mons, Belgium Jin-Sam You • Indiana Centers for Applied Protein Sciences, Indianapolis, Indiana Nina Zolotarjova • Agilent Technologies Inc., Wilmington, Delaware Kheir Zuobi-Hasona • University of Florida, Gainesville, Florida

1 Application of Fluorescence Dye Saturation Labeling for Differential Proteome Analysis of 1,000 Microdissected Cells from Pancreatic Ductal Adenocarcinoma Precursor Lesions Barbara Sitek, Bence Sipos, Günter Klöppel, Wolff Schmiegel, Stephan A. Hahn, Helmut E. Meyer, and Kai Stühler

Summary The identification of molecular changes underlying clinical pathogenic processes is often hampered by significant cellular diversity of the tissue. Pathogenic aberrant cells are surrounded by cells originating e.g., from stroma, the vascular system or other neighbouring cell types, which lead to under-representation of interesting cells when analysing whole tissue specimen. Therefore, selection of relevant cell types for detailed analysis is an absolute prerequisite for in depth elucidation of underlying biological processes. Microdissection offers the advantage to select for a biologically relevant cell type which is often in low abundance. Here, we present a proteomics approach allowing us to analyse 1,000 microdissected cells stemming from pancreatic carcinoma precursor lesions applying fluorescence dye saturation labeling.

Key Words: Difference gel electrophoresis; DIGE; fluorescence dye saturation labeling; pancreatic ductal adenocarcinoma; panin, tumor marker; two-dimensional gel electrophoresis.

1. Introduction To identify new candidate molecular markers for pancreatic ductal adenocarcinoma we established a proteomics approach analysing microdissected cells from precursor lesions, the so called pancreatic intraepithelial neoplasia From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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(PanIN) (1). Due to a limited amount of proteins available from microdissection we developed a procedure which included fluorescence dye saturation labeling in combination with high resolution two-dimensional gel electrophoresis (2-DE) (2). With this procedure we were able to analyse proteins extracted from 1,000 microdissected cells with a high resolution of up to 2,500 protein spots. For differential proteome analysis we analysed microdissected cells from 9 patients. We compared the protein expression of the different PanIN grades (PanIN 1A/B, PanIN 2, PanIN 3) and carcinoma cells related to normal epithelial cells and found 86 significantly regulated spots (p < 0.05, ratio >1.6). Using protein lysates from pancreatic carcinoma tissue as a reference proteome we were able to successfully identify the proteins after tryptic in-gel digestion. 2. Materials 2.1. Microdissection and Sample Preparation 1. 2. 3. 4. 5. 6.

Microscope: BH2 (Olympus, Wetzla, Germany). Cryostat: Cryotom SME (Shandon). Ultrasonic bath (VWR). Hand homogenisator. Injection needle: 0.65 mm × 25 mm (Braun, Melsungen, Germany). Lysis buffer: 2 M thiourea, 7 M urea, 4% 3-[(3-cholamidopropyl) dimethylammino]-1-propane sulfonate (CHAPS), 30 mM Tris-HCl, pH 8.0. 7. Hematoxylin stain: 25% (v/v) hematoxylin according to Mayer (Merck). 8. Eosin stain: 1.7% (w/v) eosin (Merck), diluted in 96% ethanol.

2.2. Cysteine-Speciﬁc Protein Labeling Using CyDye DIGE Fluor Saturation Dyes 1. Dye stock solution: CyDye DIGE Fluor saturation dyes solid compounds are reconstituted in dimethylformamide (DMF) giving a concentration of 2 mM (50 μL DMF to 100 nM of dye) for analytical gels and 20 mM (20 μL DMF to 400 nM of dye) for preparative gels. It is stable at –20°C for 3 mo. 2. 50 mM NaOH solution (for pH adjustment). 3. 2 mM TCEP (triscarboxethylphosphine) solution for analytical gels and 20 mM TCEP solution for preparative gels. 4. Image software: ImageQuantTM (GE Healthcare Bioscience). 5. Differential image analysis software: Decyder v5.02 (GE Healthcare Bioscience). 6. Fluorescence Scanner: Typhoon 9400 (GE Healthcare Bioscience).

2.3. Isoelectric Focusing 1. Seperation gel buffer: 3.5% (w/v) acrylamide, 0.3% (w/v) piperazindiacrylamide, 4% (v/v) carrier ampholines mixture (pH 2–11), 9.0 M urea, 5% glycerol, 0.06% (v/v) TEMED.

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2. Cap gel buffer: 12.3% (w/v) acrylamide, 0.13% (w/v) piperazindiacrylamide, 4% (v/v) carrier ampholyte mixture, 9.0 M urea, 5% glycerol, 0.06% (v/v) TEMED 3. 1.2% (w/v) APS (Ammoniumpersulfat). 4. Anodic buffer: 3 M urea, 7.3% (w/v) phosphoric acid. 5. Cathodic buffer: 9 M urea, 5% (w/v) glycerol, 0.75 M ethylendiamine. 6. Sephadex solution: 270 mg Sephadex suspension (20 g Sephadex was swollen in 500 mL water, resuspended into 1 L of 25% glycerol solution and filtered), plus 233 mg urea, plus 98 mg thiourea, and 25 μL ampholine mixture, pH 2–11 and 25 μL DTT (1.08 g/5 mL). 7. Protection solution: 30% urea (w/v), 5% glycerol (w/v), 2% carrier ampholytes, pH 2–4. 8. Equilibration solution: 125 mM Tris-base, 40% glycerol, 65 mM DTT, 3% SDS.

2.4. Two-dimensional Polyacrylamid Gel Electrophoresis (2D-PAGE) 1. 2. 3. 4. 5. 6. 7. 8. 9.

Glass plates compatible with fluorescence imaging (25 × 30 × 0.4 cm). Plastic spacer (30 × 1 × 0.15 cm). Apparatus for vertical SDS-PAGE. (System VA Sarstedt, Nümbrecht, Germany) Gel carrier grooves (self made). Gel solution: 570 mM Tris-base and 180 mM Tris-HCL, 0.06% TEMED, 0.2% SDS, 15% acrylamide, 0.2% bisacrylamide. Running buffer: 0.2 M Tris, 1.92 mM glycine, 0.1% (w/v) SDS. 40% (w/w) APS (Ammoniumpersulfat). Protection solution: 285 mM Tris-base and 90 mM Tris-HCL, 0.1% SDS. Agarose solution: 1% (w/w) agarose (dissolved in running buffer), 0.001% (w/w) bromphenol blue.

3. Methods The number of cells available by manual microdissection is rather limited. When analysing precursor lesions of the pancreatic adenocarcinoma only 1,000–5,000 cells can be provided in a reasonable time window (3–4 h). Due to the scarce protein amount (1,000 cells are equivalent to approx 2 μg protein) applying classical 2-DE techniques in combination with silver staining (loading amount 100 μg) or difference gel electrophoresis (DIGE) minimal labeling (50 μg) is not feasible. Therefore, DIGE saturation labeling which has a 50-fold higher sensitivity for protein detection must be applied when analysing such scarce sample amount cells (3). DIGE saturation labeling is based on covalent attachment of all protein cysteine residues prior to 2-DE (4). In contrast to DIGE minimal labeling where only one lysine residue of approx 3% of a protein species is labeled by a fluorescence dye (Cy2, Cy3 or Cy5) (5), DIGE saturation dyes leads to a complete labeling of all proteins (3).

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Moreover, in contrast to DIGE minimal labeling multiplexing within one gel for direct differential analysis is not feasible when using saturation labeling. Effects of fluorescence resonance energy transfer (FRET), by which the fluorescence emission of a CyDye can be enhanced or quenched, respectively, have to be considered (6). Therefore, only two saturation dyes are available, whereof one CyDye (mostly Cy5) is taken for differential analysis whereas the other CyDye (mostly Cy3) is taken as an internal standard for controlling system variation that ultimately provides a more accurate quantification of relative protein abundance. Furthermore, preparative protein amounts (300–500 μg) for protein identification can not be provided by microdissection. Therefore, a reference proteome stemming from a comparable source (e.g., whole tissue or cell culture lysate) must be determined. This reference proteome should match the proteome of the microdissected samples to a high degree (>90%) and thus facilitates protein identification in subsequent in-gel digestion using mass spectrometry (1). However, before commencing DIGE analysis one must optimally determine the labeling conditions i.e., the proportion of fluorescent dye to microdissected cells. It has been shown (own observations) that due to inherent differences in the cysteine content of a given proteome, coupled with variable sample conditions, one has to independently develop an optimisation procedure for each sample. This allows a high performance and high quality proteomic analysis. 3.1. Optimised Manual Microdissection for the Proteome Analysis of Pancreatic Adenocarcinoma Cells 1. Froze tissue resections containing pancreatic carcinoma on liquid nitrogen and store at –80°C (see Note 1). For histological classification of normal ducts and different PanIN stages prepare 5 μm frozen sections of peritumoral pancreatic parenchyma using a cryostat and stain with hematoxylin and eosin. 2. For microdissection in subsequent tissue slides prepare 10-μm frozen section from tissue blocks containing the required grades and only stain with haematoxylin (Fig. 1) It has been shown that eosin interferes with 2-DE leading to reduced protein recovery (Fig. 2). 3. Under microscopic observation harvest the required number of cells using a sterile injection needle. 4. Lyse the microdissected cells in 100 μL lysis buffer (4°C), sonicate (6 × 10 sec pulses on ice) after each collection step and finally centrifuge (12,000g for 5 min) the lysate and store the supernatant at –80°C.

3.2. Determination of the Minimal Number of Microdissected Cells 1. Because microdissection is very time consuming and the number of available sample material is limited it is necessary to find the minimal number of cells

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Fig. 1. Manual microdissection. Subsequent histological classification the hematoxylin stained (10 μm serial sections) PanIN lesions (A) were microdissected under a microscope using a sterile injection needle (B).

Fig. 2. Compatibility of H&E staining with 2-DE. Influence of H&E staining was investigated applying 7000 microdissected cells, each stained with H&E (A) or hematoxylin only (B). After microdissection and labeling with Cy3 the samples were separated by 2-DE and scanned using fluorescence scanner. Due to the weak protein recovery shown in (A) it is obvious that eosin interferes with proteome analysis and that hematoxylin staining is compatible with 2-DE analysis.

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Fig. 3. Determination of the minimal number of microdissected cells required for 2DE. Aliquots with 1000 and 7000 cells were labeled with Cy3 and analysed using 2-DE under the same conditions. The spots in the gels were detected using DIA mode of DeCyder software. In the gel with 1000 cells 2500 spots (A) and in the gel with 7000 cells 2600 spots (B) could be detected. sufficient for a differential proteome analysis. Therefore, a pool of approx 30,000 microdissected cells is required allowing to determine the optimal number of spots by a dilution experiment. 2. Prepare four aliquots containing 1,000, 2,500, 5,000, and 7,000 cells in the volume of 100 μL lysis buffer, respectively (see Note 2). 3. For protein labeling use the standard protocol according to user manual. Briefly, add 4 nM TCEP to each sample and incubate for 1 h at 37°C. Label the samples with 8 nM saturation dyes for 30 min at 37°C. For stopping the labeling reaction add 10 μL DTT and 10 μL Ampholytes, pH 2–4, before 2-DE. 4. Subsequent to 2-DE scan the gels as described elsewhere (see Note 3) and detect the number of spots using DIA module of DeCyder software (see Note 4). The optimal number of cells considered for subsequent analyses results from highest number of protein spots per analysed number of cells (Fig. 3).

3.3. Optimisation of Fluorescence Dye Labeling 1. Having established the optimal number of cells for comprehensive proteome coverage, one must subsequently empirically optimise the labeling conditions. This avoids any effects of over- and under-labeling. Therefore, a so called

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Fig. 4. Effects of different DIGE saturation labeling conditions detected by same/same experiment. For the optimal application of DIGE saturation labeling the labeling conditions have to be optimized. (A) If too much dye amount has been applied over-labeling leads to a horizontal shift detected in CyDye-dependent manner (arrows). (B) Under-labeling of proteins with CyDye results in vertical streaking. (C) The optimal label condition is reached if an exact overlay of both fluorescence images can be detected.

“same/same” experiment titrating different dye amounts is performed. Thus, prepare six aliquots containing the optimal number of cells (see Chapter 3.2) from the pool of 30,000 cells. 2. Label three of them with 2, 4, and 8 nM of Cy3 and another three with 2, 4, and 8 nM of Cy5. 3. Mix samples with equal dye amount and perform 2-DE for same/same experiment. 4. After gel scanning analyse the overlay images in order to find the optimal ratio of protein and dye showing an accurate overlay of Cy3 and Cy5 images using ImageQuantTM . As shown in Fig. 4, applying too much dye results in horizontal shifts could occur due to additional labeling (e.g., -amino group of lysines), whereas an insufficient amount of dye causes vertical streaks due to inadequate protein labeling.

3.4. Reference Proteome for Internal Standardization and Protein Identiﬁcation 1. 2-DE analysis of 1,000 microdissected cells (∼2 μg) does not deliver sufficient sample material for protein identification using mass spectrometry. Therefore, a reference proteome from a closely related source has to be defined allowing protein identification by protein spot assignment. Additionally, this reference proteome may be considered as an internal standard (labeled with Cy3) which reduces consumption of microdissected samples. 2. It is therefore imperative that a suitable reference proteome is identified i.e. one which not only demonstrates a high correlation to microdissected cells, but also provides an adequate protein concentration for subsequent identification.

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3.

4. 5. 6. 7. 8.

9.

Therefore freeze several tissues containing pancreatic carcinoma on liquid nitrogen directly after resection and store at –80°C. For homogenization use 100 mg of the tissue on liquid nitrogen in 148 μL lysis buffer using a hand homogeniser and sonicate the sample 6 times for 10 sec on ice. To remove insoluble debris centrifuge (12,000g for 5 min) the homogenisate. Label 3 μg of each lysate with 2 nM Cy3 according to the standard labeling protocol according to user manual. Label aliquots of 1,000 microdissected cells with 4 nM Cy5. Pair-wise mix the labeled tissue lysates with labeled cells and process by 2-DE. After scanning, analyze the images using DIA module of DeCyder (Fig. 5) and calculate a matching rate between microdissected cells and carcinoma tissue (see Note 5). Optimize the labeling conditions for the tissue lysate showing the highest matching rate with the microdissected cells also by same/same experiment (see Chapter 3.3).

Fig. 5. Protein spot pattern of carcinoma tissue (reference proteome) and microdissected cells after 2-DE. For protein identification and internal standardization a reference proteome with high degree in protein spot matching is necessary. For the proteome of a pancreatic tissue lysate a high matching rate (>90%) has been determined. In section A all detected protein spots of carcinoma tissue and microdissected cells, respectively, can be matched (letters are shown for better assignment). Whereas in section B beside a number of assignable protein spots (numbers) a protein spot unique for microdissected cells (arrows) was revealed.

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3.5. Two-Dimensional Gel Electrophoresis 1. These instructions are modified from the 2-DE technique as described by Klose and Kobalz (2). This technique is based on isoelectric focusing employing carrier ampholyte tube gels. It can be easily adapted to the DIGE technique and other formats, including analytical as well as preparative gels. In spite of the fact that most of the instruments were constructed in-house, equivalent equipment is commercially available. For the second dimension the Desaga VA300 gel system is applied. 2. Two days before running isoelectric focusing (IEF) tube gels (Ø 1.5 mm, 20 cm) are prepared. Add 45 μL of APS solution to 2 mL of separations gel solution and fill tube to first mark (20 cm) using a syringe. Now, add 14 μL APS solution to 0.7 mL cap gel solution and cast cap gel to second mark (20.5 cm) behind the separation gel. To prevent urea crystallisation place an air cushion under the cap gel to third mark (21 cm). 3. Before starting IEF apply 2 mm sephadex solution to prevent protein precipitation onto the separation gel. Then load the sample (dye-labeled) and overlay the sample with protection solution (approx 5 mm) to prevent direct contact of the acidic cathodic buffer (see Note 6). 4. Fill anodic (bottom) and cathodic buffer (top) into the IEF chambers. Ensure that no air bubbles hamper IEF (see Note 6). Start isoelectric focusing applying a step-wise voltage program (100V for 1 h, 200 V/1 h, 400 V/17.5 h, 650 V/1 h, 1,000 V/30 min, 1,500 V/10 min, 2,000 V/5 min). 5. While the 21.5 h IEF is running, gels for the second dimension should be prepared. Clean the glass plates thrice (gel side) using a lint-free paper towel— for the first wash use doubled distilled water followed by 100% ethanol and finally 70% ethanol. To ensure correct gel dimension two 1.5 mm plastic spacers are placed between two plates sealed by silicon. 6. Add 288 μL APS solution into 144 mL gel buffer, cast the gel and overlay with water-saturated isobutanol. 7. After polymerization (45 min) remove isobutanol and wash the surface with a protection solution. To protect gel drying place 2-DE protection solution onto the gel and store the gels at 4°C. 8. After IEF extrude the gel by means of inserting a nylon fiber (see Note 7) into the gel groove of the IEF gel carrier and incubate with equilibration solution for 15 min to load proteins with SDS. 9. Wash the gel three times with running buffer before applying to the second dimension. 10. For the transfer of the IEF gel into SDS-PAGE gel hold the groove with gel in contact with the edge of the glass plate and slide the gel between the glass plates using a wire suitably formed. 11. Overlay the IEF gels with agarose solution, add the running buffer to the upper and lower (15°C) chambers and start the electrophoresis. For the entrance of the proteins into the SDS-PAGE gel apply low current (75 mA) for 15min. When the proteins have entered increase the current to 200 mA for approximately 5–7 h.

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3.6. Differential Proteome Analysis of 1000 Microdissected Cells from Different PanIN Stages 1. This instruction comprised all steps obtained during optimisation procedure as described above. 2. Harvest 1000 cells by manual microdissection and incubate the cells in 100 μL lysis buffer. 3. For protein labeling according to the optimised protocol reduce the proteins with 2 nM TCEP and label the proteins of the microdissected cells with 4 nM Cy5. 4. For the preparation of the internal standard label 3 μg protein from a PDAC tissue sample lysate with 2 nM Cy3. 5. Add 10 μL DTT to stop labeling reaction and add 10 μL Ampholytes, pH 2–4.

Fig. 6. Representative images for each investigated progression step of PDAC. The lysates from 1000 microdissected cells were labeled with Cy5 mixed with internal standard and processed by 2-DE. Subsequent image acquisition the spot patterns were analyzed using DeCyder software and 2,000–2,500 protein spots have been detected for the different PanIN grades. Protein spot 2574 has been detected as differentially expressed in PanIN 2, PanIN 3 and Carcinoma (see Fig.7)

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6. Mix the sample and analyse the mixture using 2-DE. 7. After 2-DE, leave the gels between the glass plates and acquire the images using the Typhoon 9400 scanner (see Note 3). Therefore, choose excitation wavelengths and emission filters specific for each of the CyDyes according to the Typhoon user guide. 8. Before image analysis with DeCyder software crop the images with ImageQuantTM software (Fig. 6). 9. For intra-gel spot detection and quantification use the Differential In-gel Analysis (DIA) mode of the DeCyder software. Set the estimated number of spots to 3000. Apply an exclusion filter to remove spots with a slope greater than 1.6 (see Note 4). 10. After spot matching between the different gels using the Biological Variation Analysis mode (BVA) consider only protein spots with an expression changes of factor > 1.6 and p-value (Student’s t-test) < 0.05 as significantly regulated (Fig. 7).

Fig. 7. Stage-dependent regulation of protein spot 2574. For each patient and PanIN stage the protein spot intensity is shown. The depicted protein spot 2574 (see Fig. 6) shows a significant up-regulation (p < 0.05) in the carcinoma stage by a factor of 3.2. In the PanIN stages 2 and 3 a significant down-regulation by a factor of -2.7 and -1.8 has been determined.

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3.7. Micropreparation of Signiﬁcantly Regulated Protein Spots 1. After differential analysis protein spots are identified using mass spectrometry (MALDI-MS, ESI-MS) subsequent tryptic in-gel digestion. Therefore, 100-fold protein amount of the internal standard (reference proteome) must be labeled and applied to 2-DE. A preparative label kit with 400 nM Cy3 is available. 2. For preparative gels label 400 μg of tissue lysate with 260 nM of Cy3 (130 nM TCEP) (see Note 8). 3. Directly after gel scanning isolate the protein spots of interest manually (see Note 9). Assign the positions of the spots using a printout of the gel placed underneath the glass plate. 4. Put the isolated protein spots into sample cups (glass) and store them at –80°C. 5. For protein identification different mass spectrometric (MS) techniques can be applied subsequent to enzymatic in-gel digestion. Matrix assisted laser desorption/ionization MS (MALDI-MS) allows a fast and sensitive MS analysis performing peptide mass fingerprinting (PMF) (7). In cases where more sequence information is necessary (e.g. posttranslational modification) liquid chromatography-coupled electrospray ionisation MS (LC-ESI-MS) with its high performance for peptide fragment mass fingerprinting (PFF) is preferable (8).

4. Notes 1. To protect samples against tissue disruption (freezer burn) after resection the tissue is placed in tin foil at first and then stored in a cryotube. 2. Usually, depending of tube’s diameter not more than 50 μL can be applied. Therefore, we increased the volume by blowing up the glass tube so that a sample volume of 100–200 μL can be applied. 3. Before scanning the glass plates have to be cleaned thoroughly. First of all use ethanol for removing of acrylamide or silicone and then clean the glass plates with water. 4. To avoid the detection of dust particles and artifacts as protein spots in the gel, an exclusion filter concerning slope, area, peak height, or volume can be applied. For one or more of these characteristics a value that distinguishes dust particles and spots has to be found. 5. For the determination of the matching rate between two images which derived from the same gel analyze the gel using DIA mode of DeCyder software. After a spot detection an average ratio of 2.0 could be set in order to find spots occurring in both images (Average ratio < 2.0). 6. Air bubbles should be avoided by applying the solution slowly under the surface (1 mm) of the solution which has been applied before. For Sephadex application the small volume of separation gel buffer generated during polymerisation is sufficient. 7. To prevent destruction of the IEF gel by the nylon fiber (i) the thermoplastic nylon fiber should be fitted to the tube inner diameter by melting one end into the tube and (ii) the gel should be extruded using the cap gel (acrylamide concentration

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12.3%) as a cushion or (iii) another possibility is to polymerise a high concentrated acrylamide solution (15%) above the extruding side and use this gel piece as a cushion. 8. To avoid a high sample volume for the preparative labeling TCEP and Cy3 should have a concentration of 20 mM respectively instead of 2 mM (see Chapter 2.2). The amount of TCEP and Cy3 for labeling of preparative protein lysate has to be calculated according to labeling conditions for analytical gels. 9. Before scanning, mark the glass plate from the picking gel with fluorescence stickers The stickers are necessary for matching between the gel (the proteins are not visible) and the gel image. After scanning print the gel image in the original size. Put the print under the glass plates, align the position of the stickers on the glass plate with the image.

Acknowledgments The authors would like to thank Kathy Pfeiffer, Conny Bieling, Sabine Burkert and Birgit Streletzki for excellent technical assistance and Jon Barbour for critical reading of the manuscript. This work was supported by the grant from the Deutsche Krebshilfe (B.S., J.L., S.A.H and K.S., 70-2988-Schm3), Bundesministerium für Bildung und Forschung (NGFN, FZ 031U119) and the Nordrhein Westfalen Ministerium für Wissenschaft und Forschung.

References 1. Sitek, B., Lüttges, J., Marcus, K., Klöppel, G., Schmiegel, W., Meyer, H. E., Hahn, S. A. and Stühler, K. (2005) Application of fluorescence difference gel electrophoresis saturation labeling for the analysis of microdissected precursor lesions of pancreatic ductal adenocarcinoma. J. Proteomics, 5, 2665–2679. 2. Klose, J. and Kobalz, U. (1995) Two-dimensional electrophoresis of proteins: an updated protocol and implications for a functional analysis of the genome. J. Electrophoresis 6, 1034–1059. 3. Kondo, T., Seike, M., Mori, Y., Fujii,K., Yamada, T., and Hirohashi, S. (2003) Application of sensitive fluorescent dyes in linkage of laser microdissection and two-dimensional gel J. Proteomics 3, 1758–1766. 4. Sitek, B., Scheibe, B., Jung, K, Schramm ,A., and Stühler, K. (2006) Difference gel electrophoresis (dige): the next generation of two-dimensional gel electrophoresis for clinical research, in Proteomics in Drug Research (Hamacher et al., eds), WileyVCH, Weinberg, pp 33–55. 5. Unlü, M., Morgan,M. E., and Minden,J. S. (1997) Difference gel electrophoresis: a single gel method for detecting changes in protein extracts. Electrophoresis 18, 2071–2077. 6. Gruber, H. J., Hahn, C. D., Kada, G., Riener, C. K., Harms, G. S., Ahrer, W., Dax, T. G., and Knaus, H.-G. (2000) Anomalous fluorescence enhancement of Cy3

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and cy3.5 versus anomalous fluorescence loss of Cy5 and Cy7 upon covalent linking to IgG and noncovalent binding to avidin. Bioconjug. Chem. 11, 696–704. 7. Stühler, K., and Meyer, H. E. (2004) MALDI: more than peptide mass fingerprints. Curr Opin Mol Ther. 3, 239–248. 8. Marcus, K., Moebius, J., and Meyer, H. E. (2003) Differential analysis of phosphorylated proteins in resting and thrombin-stimulated human platelets. Anal Bioanal Chem. 7, 973–993.

2 Albumin and Immunoglobulin Depletion of Human Plasma Rosalind E. Jenkins, Neil R. Kitteringham, Carrie Greenough, and B. Kevin Park

Summary Plasma and serum have been the focus of intense study in recent years in the expectation that they will provide important biomarkers of health and disease, without the need for invasive procedures. This aim has been hindered by the fact that a few highly abundant proteins dominate the protein profile, masking the lower abundance proteins and limiting our ability to analyse the entire plasma proteome. This chapter details a simple and effective method for removal of two of the most dominant proteins in plasma, albumin and immunoglobulin.

Key Words: Affinity chromatography; albumin; depletion; immunoglobulin; plasma proteome.

1. Introduction The dynamic range of proteins in human plasma is greater than ten orders of magnitude, from albumin present at around 30 mg/mL to cytokines such as interleukin 6 present at basal levels of 1–3 pg/mL (1). This extraordinarily wide range of protein concentrations within a single compartment makes detailed analysis of the lower abundance proteins technically demanding, yet it is exactly these low level species that are likely to provide insight into disease states, into the effects of therapeutic intervention, and to yield specific and sensitive biomarkers. Many methods have been developed to deplete serum and From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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plasma of the high abundance, housekeeping proteins, including alcohol precipitation, ultracentrifugation, salting in/salting out (2,3), and molecular weight fractionation (4,5). However, removal of proteins by affinity capture is the most commonly used approach because of the highly selective nature of the depletion. Anti-human albumin monoclonal antibody immobilized onto a solid support such as sepharose forms the basis of many immunoaffinity columns for the removal of the most abundant protein in human plasma. Clearly the affinity and specificity of the antibody determine the efficiency and discrimination of protein removal. However, the support on which the antibody is immobilized will also impact on the process, not least in terms of the binding capacity of the matrix. Affinity matrices based on bacterial protein A, protein G, or protein L, or molecularly engineered versions of these, are most commonly used to isolate immunoglobulins (6–11). Sophisticated multicomponent immunoaffinity matrices that are designed to deplete 10–15 of the most abundant plasma proteins are available (12) and have been used with great success, but they do have several drawbacks: they are expensive; they have a relatively low binding capacity so that several aliquots of plasma must be processed individually and then pooled to generate sufficient material for further analysis; and the depletion of the target proteins is frequently far from complete. The method described in this chapter, although limited to the depletion of only two of the highest abundance proteins, is simple, rapid, and accessible and provides a significant improvement in coverage of the plasma proteome obtainable by 2D gel electrophoresis. 2. Materials 2.1. Blood Sampling 1. Heparinized tubes. 2. Refrigerated centrifuge (up to 450g). 3. 0.5-mL Eppendorf tubes.

2.2. Albumin Depletion 1. 2-mL cartridge containing POROS® beads coated with monoclonal goat antihuman serum albumin (HSA) (Applied Biosystems). 2. Loading and washing buffer: phosphate buffered saline (PBS): 137 mM NaCl, 1 mM KH2 PO4 , 5.6 mM Na2 HPO4 , 2.7 mM KCl, pH 7.4. 3. Elution buffer: 12 mM HCl. 4. HSA column wash buffer: 1 M NaCl. 5. High performance liquid chromatography system capable of flow rates from 0.5–3 mL/min (for automated depletions).

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2-mL and 10-mL plastic syringes (for manual depletions). Blunt ended syringe needle (for manual depletions). 1.5-mL Eppendorf tubes. Spectrophotometer able to read the absorbance at 280 nm (for manual depletions).

2.3. Immunoglobulin Depletion 1. 0.2 mL cartridge containing POROS® beads coated with recombinant protein G (Applied Biosystems). 2. Loading and washing buffer: phosphate buffered saline (PBS). 3. Elution buffer: 12 mM HCl. 4. Protein G column wash buffer: 1 M NaCl 10% acetic acid. 5. High performance liquid chromatography system capable of flow rates from 0.5–3 mL/min (for automated depletions). 6. 2-mL and 10-mL plastic syringes (for manual depletions). 7. Blunt ended syringe needle (for manual depletions). 8. 1.5-mL Eppendorf tubes. 9. Spectrophotometer able to read the absorbance at 280 nm (for manual depletions).

2.4. One-Dimensional (1D) gel Electrophoresis 1. Standard 1D gel electrophoresis apparatus. 2. Laemmli sample buffer (1×): 3% SDS, 0.1 M Tris-HCl, 15% glycerol, 0.2% bromophenol blue, pH 7.6.

2.5. TCA Precipitation of Proteins 1. 20% trichloroacetic acid (TCA) and ice-cold acetone. 2. Resuspension buffer: 5% SDS, 1.15% DTT. 3. Refrigerated centrifuge (up to 14,000g).

2.6. 2D Gel Electrophoresis 1. Standard 2D gel electrophoresis apparatus. 2. IPG rehydration buffer: 9 M urea, 2% (w/v) CHAPS, bromophenol blue (trace), 2% IPG buffer (Pharmacia), 0.28% DTT

3. Methods 3.1. Collection of Samples 1. Collect the blood into heparinized tubes and sediment the red blood cells as soon after acquisition as possible by centrifugation at 450g for 10 min (see Notes 1, 2, and 3).

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2. Recover the supernatant, divide it into small aliquots and store them at –80°C. Each aliquot should be thawed and used only once. 3. Once thawed, centrifuge the aliquot briefly to remove any precipitate that may have formed at low temperatures and that may block the affinity cartridges.

3.2. Albumin Depletion 1. Samples are diluted to 6 mg/mL in PBS and 600 μL (equivalent to 3.6 mg protein or approx 60 μL plasma) are applied to the column (see Notes 4, 5 and 6). 2. The POROS column is mounted in a cartridge holder supplied by the manufacturer with ports for insertion of standard HPLC fittings, if performing the following steps robotically, or for a needle port adapter at the top and a short piece of PEEK tubing at the base to help direct the flow-through when performing the steps manually. The blunt-ended needle attached to a 10-mL syringe is inserted firmly into the needle port adapter for manual application of the sample and buffers to the column, which should be applied at a flow rate of one drop per second throughout for manual chromatography. The flow rates for depletions performed on an HPLC system are given at the appropriate points in the text. 3. The column is equilibrated by applying 10 column volumes (20 mL) of PBS at a flow rate of 2.4 mL/min. 4. The diluted plasma sample is applied to the column at a flow rate of 1.2 mL/min and the flow-through (containing plasma proteins minus albumin) is collected as 500-μL fractions into 1.5-mL eppendorf tubes. 5. A further 10 column volumes (20 mL) PBS are applied to the column to ensure that all non-specifically bound proteins are flushed through, with continued collection of fractions. Collection of a total of ten fractions is generally sufficient to capture all of the nonbound plasma proteins. 6. Albumin is eluted from the column by the application of 5 column volumes (10 mL) of 12 mM HCl at a flow rate of 2.4 mL/min, with the first five 1-mL fractions collected into 1.5-mL eppendorf tubes. 7. The column is cleaned by applying 10 column volumes (20 mL) of HSA column wash buffer (1 M NaCl) at a flow rate of 2.4 mL/min (see Note7). 8. If further samples are to be processed, the column may be equilibrated with PBS as in point 3. If the column is to be stored for 1–2 d, flushing through with 10 column volumes (20 mL) PBS and storage at 4°C will be sufficient. If it is to be stored for a longer period, the PBS should contain 0.05% sodium azide.

3.3. Assessment of Fractions Containing Albumin-Depleted Plasma Proteins by 1D Gel Electrophoresis (see Note 8) 1. Aliquots of 10 μL of the flow-through fractions are mixed with 4× Laemmli sample buffer, boiled and loaded onto SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) minigels. 2. Similarly, 3–6 μL of the eluted HSA fractions, and 2 μL of the dilute undepleted plasma sample, are prepared for gel analysis.

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Fig. 1. Depletion of albumin from human plasma. Coomassie blue (A) and silver (B) stained 1D gel of undepleted plasma (P), aliquots of the flow-through fractions 1–10 and eluted HSA (E1). The undepleted plasma appears to contain a relatively complex mixture of proteins that is dominated by the albumin band at 66 kDa. The depleted fractions should contain the same range of proteins but the albumin band should be completely absent. In contrast, the fraction representing eluted albumin should contain no other protein bands, even at the level of sensitivity of silver stain. The multiple bands visible below the major HSA band were determined by mass spectrometry to be breakdown products, presumably because of the acid conditions under which the albumin was eluted from the column. (C) UV trace of albumin depletion performed by HPLC showing that fractions 3–7 contain the protein flow-through from the anti-HSA column whereas fractions 17 and 18 contain the eluted HSA (E1) itself. 3. The samples are loaded onto 10% SDS-PAGE minigels and the gels stained with colloidal Coomassie Blue (Fig. 1A) or silver stain (Fig. 1B) after electrophoresis.

3.4. Assessment of Fractions Containing Albumin-Depleted Plasma Proteins by UV Absorbance 1. Once the depletion method has been optimized, the flow-through fractions may be assessed by spectrophotometry. If samples have been depleted using an HPLC system, a real-time UV trace of the chromatographic separation is usually available. Fig. 1 is an example of such a trace.

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2. If performing the depletions manually, the fractions may be assessed using an off-line spectrophotometer. Aliquots of 100 μL of each fraction are placed in mini-cuvets or wells of a 96-well plate, and the absorbance at 280 nm recorded. 3. All flow-through fractions producing an absorbance value are pooled for the next step in the protocol, the depletion of immunoglobulin (see Note 9).

3.5. Immunoglobulin Depletion 1. The 0.2-mL protein G POROS column is supplied with a smaller cartridge holder and should be mounted as described earlier. The blunt-ended needle attached to a 2-mL syringe is used for manual application of sample and buffers to the column. As for the HSA depletion column, all reagents are applied to the column at a flow rate of 1 drop per second when the chromatography is performed manually. Flow rates for depletion using an HPLC system are noted at appropriate points in the text. 2. The column is equilibrated by applying 10 column volumes (2 mL) of PBS at a flow rate of 1mL/min. 3. The albumin-depleted plasma sample cannot be applied as a single aliquot but should be split into two aliquots of 600 μL (see Note 10), each being loaded at a flow rate of 0.5 mL/min. The flow-through (containing plasma proteins minus albumin and immunoglobulin) is collected as 500-μL fractions into 1.5-mL eppendorf tubes. 4. A further 10 column volumes (2 mL) PBS are applied to the column at a flow rate of 1 mL/min to ensure that all nonspecifically bound proteins are flushed through, with continued collection of fractions. Collection of a total of six fractions is generally sufficient to capture all the nonbound plasma proteins. 5. Immunoglobulin is eluted from the column by the application of 10 column volumes (2 mL) of 12 mM HCl at a flow rate of 1 mL/min, with two 1-mL fractions being collected into 1.5-mL eppendorf tubes. 6. After the first aliquot has been depleted of immunoglobulin, the column is reequilibrated with PBS as in point 2, and the second aliquot processed as in steps 3–5. 7. After the second aliquot has been processed, 10 column volumes (2 mL) of protein G column wash buffer (1 M NaCl containing 10% acetic acid) are applied to the column at a flow rate of 1 mL/min (see Note 11). As for the anti-HSA column, the protein G column may be stored for 1–2 days with PBS but for longer periods, the PBS should contain 0.05% sodium azide.

3.6. Assessment of Fractions Containing Albuminand Immunoglobulin-Depleted Plasma Proteins by 1D Gel Electrophoresis (see Note 8) 1. Aliquots of 10 μL of the flow-through fractions are mixed with 4× Laemmli sample buffer, boiled and loaded onto SDS-PAGE minigels.

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Fig. 2. Depletion of immunoglobulin from albumin-depleted human plasma. Coomassie blue (A) and silver (B) stained 1D gel of undepleted plasma (P), HSA-depleted plasma (H), aliquots of the flow-through fractions 1–6 and eluted immunoglobulin (E2). The depletion of the immunoglobulin bands is more difficult to discern than the depletion of the very abundant albumin band shown in figure 1, but the eluted protein should be clearly seen as two bands representing the heavy and light chains of the immunoglobulin (IgH and IgL ). (C) UV trace of immunoglobulin depletion performed by HPLC showing that fractions 1 and 2 contain the protein flow-through from the protein G column whereas fraction 7 contains the eluted immunoglobulin.

2. Similarly, 3–6 μL of the eluted immunoglobulin fractions, 2 μL of the dilute undepleted plasma, and 10 μL of the albumin-depleted plasma are prepared for gel analysis. 3. The samples are electrophoresed on 10% SDS-PAGE minigels and the gels stained with colloidal Coomassie Blue or silver stain (Fig. 2).

3.7. Assessment of Fractions Containing Albumin- and Immunoglobulin-Depleted Plasma Proteins by UV Absorbance 1. Once the depletion method has been optimized, the flow-through fractions may be assessed by simple spectrophotometry by measuring their absorbance in-line

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at 214 nm using the UV detector on the HPLC system, or off-line at 280 nm in a cuvet or microplate format. 2. All flow-through fractions producing an absorbance value are pooled before processing for 2D gel electrophoresis or other proteomic analysis (see Note 12).

3.8. TCA Precipitation of Depleted Protein Samples for 2D Gel Electrophoresis 1. The depleted samples should be maintained at 4° C or on ice throughout this procedure. 2. A solution of 20% TCA in water is prepared just before use and placed on ice to chill (see Note 13). 3. 2 mL 20% TCA are added to 2 mL of the dilute depleted plasma, and the sample is mixed gently and incubated on ice for 30 min. 4. The sample is then divided equally between three 1.5-mL eppendorf tubes and centrifuged at 14,000g and 4° C for 10 min to pellet the proteins.

Fig. 3. 2D gel images of undepleted (A) and depleted (B) plasma. The plasma proteins were focussed on nonlinear pH 3–10 IPG strips before 2nd dimension separation on 12% SDS PAGE gels. The gels were stained with colloidal Coomassie blue. Image analysis of the gels shows that the albumin and immunoglobulin heavy chain are 98% and 80% depleted, respectively. The resolution of the protein spots is improved significantly by depletion, particularly for proteins migrating in the top half of the gel close to the migration position of albumin. Indeed, peptide mass fingerprinting of the visible features reveals that hemopexin comigrates with albumin, yet it is undetectable on gels of the undepleted sample. There is an overall increase in the number of detectable spots of approx 50% when the gels are stained with Coomassie blue, indicating that the detection of lower abundance proteins has been improved.

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5. The supernatant is discarded, and the pellets washed with 2 × 1 mL of ice-cold acetone to remove all traces of the TCA. 6. The pellets are air-dried briefly (approx 5 min) before resuspension in 12 μL resuspension buffer (see Note 14). 7. The sample is heated at 95° C for 10 min to aid solubilisation followed by the addition of 350 μL IPG-strip rehydration buffer. 8. The sample is clarified by a high speed centrifugation step before conventional 2D electrophoresis (Fig. 3).

4. Notes 1. To make meaningful comparisons between plasma samples, it is vital that the collection and storage is consistent. Plasma is rich in proteases and inappropriate or overlong storage can lead to significant changes in the protein profile. Whatever procedure is chosen for sample preparation, it must be strictly adhered to. 2. Serum and plasma are almost identical in terms of the protein profile observed on 2D gels. However, some of the proteins involved in the clotting process, such as fibrinogen, are removed when the clotted red blood cells are separated from the serum by centrifugation. There is also a slightly increased tendency for lysis of the red blood cells when preparing serum. It is probably easier and more consistent to prepare plasma. 3. The anti-HSA column may be used to deplete albumin from the plasma or serum of various animal species, but lower levels of total protein must be loaded onto the column. We have depleted albumin from mouse and rat serum after loading 1–2 mg total protein. The protein G column works as effectively for animal immunoglobulins as for human. 4. Most human plasma samples have a protein concentration of 60–70 mg/mL with approximately half of that comprised of albumin. The binding capacity of a 2 mL anti-HSA POROS cartridge is 1.8–2 mg albumin so the samples will require a dilution step before loading onto the column. 5. Both the anti-HSA and the protein G POROS columns may be used for the depletion of multiple samples without loss of effectiveness, as long as they are cleaned and stored correctly. The smaller protein G column does have a tendency to deteriorate before the larger anti-HSA column, but we have successfully processed at least 50 plasma samples through one pair of columns. 6. Manual depletions are just as effective as those performed on HPLC systems, but they are rather tedious. A cheap but effective alternative is to use a syringe pump to apply samples and buffers. 7. The anti-HSA column should be cleaned with wash buffer (1 M NaCl) after every depletion to prevent residual protein build-up that would lead to decreased efficiency of albumin removal and increased pressure during the chromatography steps.

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Jenkins et al.

8. Measuring the absorbance at 214/280 nm provides a measure of the total amount of protein in each fraction, but it does not indicate how much of the target protein has been removed (efficiency of depletion), nor whether other proteins have been depleted as well (specificity of depletion). A visual assessment of these factors can be rapidly achieved by 1D gel electrophoresis. However, once the protocol has been optimized, it should be necessary to perform this sort of quality control only if the UV trace looks anomalous, or when a new POROS cartridge is being employed. 9. At this stage, it is usual to have a total volume of approx 1.5 mL of albumindepleted plasma containing roughly 900 μg of protein. Because the amount of protein loaded onto the column was approx 3.6 mg, the reduction in total protein content is equivalent to 75% suggesting that the depletion of albumin is complete. 10. Immunoglobulin comprises 8–26% of the total protein in plasma and is therefore present at concentrations of 5–18 mg/mL. The sample has already been diluted at least 1:10 during the albumin depletion, so the concentration is now approx 0.5–1.8 mg/mL. The binding capacity of the protein G cartridge is up to 1.8 mg immunoglobulin, so the maximum volume of the pooled fractions containing HSA-depleted plasma that could be loaded is 1 mL. These fractions may be stored at 4° C for short periods of time before the immunoglobulin depletion, but should be processed within 8h. 11. The protein G column should be cleaned after 2 immunoglobulin depletions to keep it functioning at the highest efficiency. After the first depletion, the column is simply re-equilibrated with PBS as in Section 2.3.5 point 1, but after the second, it should be exposed to wash buffer (1 M NaCl/10% acetic acid). 12. At this stage, it is usual to have a total volume of approx 2 mL of depleted plasma containing roughly 500 μg protein. This is equivalent to a further 11% reduction in protein content of the plasma, which is approximately what would be expected following removal of the immunoglobulin. However, this is too dilute for most proteomic analyses so a method to concentrate the proteins must be employed. 13. There are several reagents for precipitating proteins from dilute samples, including acetone, methanol and acetonitrile, but the method that seems to work most effectively for the samples described here is TCA precipitation. It is essential that the TCA is prepared immediately before use: even storage for 2–3 h reduces the efficiency of precipitation. The sample may be precipitated with TCA for longer than 30 min, but there is a risk of protein degradation or modification on prolonged exposure. Alternative methods for protein concentration include molecular weight cut-off filters, for which a significant loss of total protein is a factor, or the use of reversed phase matrices to capture the proteins, for which the elution buffer must be very carefully selected for its compatibility with the 1st dimension separation. 14. The volume of resuspension buffer (5% SDS, 1.15% DTT) to be added must be calculated carefully to ensure that after mixing with IPG-strip rehydration buffer, the level of SDS is within tolerable levels for the 1st dimension

Albumin and Immunoglobulin Depletion of Human Plasma

25

separation, i.e., less than 0.25%. When the sample described here is mixed with 350-μL IPG-strip rehydration buffer, the final concentration of SDS in the buffer is 0.17%, well within the tolerance of the 1st dimension separation.

Acknowledgments This work was supported by the Wellcome Trust. Thanks to Rod Watson of Applied Biosystems for assistance with the establishment of protocols. References 1. Anderson, N. L., and Anderson, N. G. (2002) The human plasma proteome: history, character, and diagnostic prospects. Mol. Cell Proteomics 1, 845–67. 2. Cohn, E. J. (1941) The properties and functions of the plasma proteins with a consideration of the methods for their separation and purification Chem. Rev. 28, 395. 3. Simoni, R. D., Hill, R. L., and Vaughan, M. (2002) The beginning of protein physical chemistry. Determinations of protein molecular weights. The work of Edwin Joseph Cohn. J. Biol. Chem. 277, 19e. 4. Georgiou, H. M., Rice, G. E., and Baker, M. S. (2001) Proteomic analysis of human plasma: Failure of centrifugal ultrafiltration to remove albumin and other high molecular weight proteins Proteomics 1, 1503–06. 5. Tirumalai, R. S., Chan, K. C., Prieto, D. A., Issaq, H. J., Conrads, T. P., and Veenstra, T. D. (2003) Characterization of the low molecular weight human serum proteome Mol. Cell Proteomics 13, 13. 6. Akerstrom, B., Brodin, T., Reis, K., and Bjorck, L. (1985) Protein G: a powerful tool for binding and detection of monoclonal and polyclonal antibodies J. Immunol. 135, 2589–92. 7. Forsgren, A., and Sjoquist, J. (1966) “Protein A” from S. aureus. I. Pseudo-immune reaction with human gamma-globulin J. Immunol.97, 822–7. 8. Housden, N. G., Harrison, S., Roberts, S. E., Beckingham, J. A., Graille, M., Stura, E., and Gore, M. G. (2003) Immunoglobulin-binding domains: Protein L from Peptostreptococcus magnus Biochem. Soc. Trans. 31, 716–8. 9. Kabir, S. (2002) Immunoglobulin purification by affinity chromatography using protein A mimetic ligands prepared by combinatorial chemical synthesis Immunol. Invest. 31, 263–78. 10. Svensson, H. G., Hoogenboom, H. R., and Sjobring, U. (1998) Protein LA, a novel hybrid protein with unique single-chain Fv antibody- and Fab-binding properties Eur. J. Biochem. 258, 890–6. 11. Wilchek, M., Miron, T., and Kohn, J. (1984) Affinity chromatography Methods Enzymol. 104, 3–55. 12. Pieper, R., Su, Q., Gatlin, C. L., Huang, S. T., Anderson, N. L., and Steiner, S. (2003) Multi-component immunoaffinity subtraction chromatography: An innovative step towards a comprehensive survey of the human plasma proteome Proteomics 3, 422–32.

3 Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase Chromatography of Human Serum James Martosella and Nina Zolotarjova

Summary Serum analysis represents an extreme challenge because of the dynamic range of the proteins of interest, and the high structural complexity of the constituent proteins. High-abundant proteins such as albumin, IgG, transferrin, haptoglobin, IgA and alpha1anti-trypsin represent up to 85% of the total protein mass in serum (Fig. 1). These major protein constituents interfere with identification and characterization of important moderate- and low-abundant proteins by limiting the dynamic range of mass spectral and electrophoretic analysis. During protein isolation, separation, and analysis, these six proteins often mask the detection of the more important low-abundant proteins that are of high interest as biomarkers of disease or drug targets. In one- and two-dimensional gel electrophoresis (1DGE and 2DGE) for example, the spots or bands because of these six highly abundant proteins, as well as their fragments, often overlap or completely mask large regions of the gel, making detection of the myriad low-abundant proteins very difficult, if not impossible. Moreover, proteomic analysis methods commonly include an electrophoretic or chromatographic separation step which, of course, has a finite mass loading tolerance. The presence of a large quantity of high-abundant proteins limits the mass load of targeted proteins that can be initially sampled by these separation methods and thus requires the need for multidimensional separation techniques to reduce sample complexity. Herein we describe immunoaffinity depletion combined with reversed-phase separation modes to reduce the sample complexity of human serum. We selectively immunodepleted six of the most abundant proteins from human serum, then employed gradient elution reversed-phase (RP) HPLC to fractionate the remaining serum proteins. The workflow shown in (Fig. 2) was optimized to process immunodepleted flow-through serum samples directly to a RP column with minimal sample handling. The RP operational conditions permitted robust and repeatable separations and have been optimized specifically for immunodepleted serum samples. From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

27

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Martosella and Zolotarjova

Key Words: affinity chromatography; human plasma; human serum; HPLC; immunodepletion; prefractionation; proteomics; reversed-phase chromatography.

Alpha-1-antitrypsin 3.8% Immunoglobulin A 3.4% Transferrin 3.3%

Immunoglobulin G 16.6%

Haptoglobin 2.9%

Other 15%

Albumin 54.3%

Fig. 1. Composition of proteins in human serum. The protein composition is schematically depicted based on mass abundance in normal human serum. The six high-abundant proteins removed by the immunoaffinity column comprise approx 85% of the total protein mass in human serum.

Human Serum or Plasma

Immunodepletion of Six High-abundant Proteins Sample Denaturation

High Temp. RP-HPLC 3-Step Multi-Segment Gradient

Multiple RP HPLC Fraction Collection

2D-PAGE LC/MS/MS

Fig. 2. Multidimensional chromatographic workflow.

Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase

29

1. Introduction Interest in proteomic analysis of human serum has been greatly elevated during the past several years as liquid chromatography (LC) and mass spectrometry (MS) methodologies have evolved sufficiently to investigate this challenging sample. The popularization of multidimensional LC methods, and the ever-improving sensitivity and performance of multi-stage MS instruments, combined with highspeed database searching, is permitting complex protein samples to undergo analysis by identification of constituent tryptic peptide fragments. Proteomic analysis of human serum represents an extreme challenge because of the dynamic range of the proteins of interest. Serum (plasma) contains many proteins, estimated to include almost 500,000 molecular species and has more than 10 orders of magnitude concentration range, from mg/mL to pg/mL, and possibly less (1). This wide range of analytical target molecules is currently outside the realm of the dynamic range of available technologies for proteomic analyses. A means to address the complexity of these samples is the application of multidimensional separation techniques (2,3), for example, by multidimensional LC fractionation. We describe an immunoaffinity and RP LC column approach, which reduces the complexity of serum or plasma. Samples are first immunodepleted of the six most abundant proteins using an immunoaffinity LC method. This separation delivers a flow-through fraction containing low-abundant proteins, whereas the bound high-abundant proteins are left behind. Second, the immunodepleted samples are separated under a set of optimized reversed-phase (RP) conditions using a highrecovery macroporous column material. The conditions and protocol have been designed specifically for immunodepleted human serum or plasma samples. The combination of sample simplification by immunoaffinity depletion, combined with robust and high recovery RP-HPLC fractionation, yields samples permitting higher quality protein identifications (4). The approach presented here enables an expanded dynamic range for the detection of low-abundant proteins in the complex proteomic samples and thereby assists in the search for novel biomarkers of disease states and intervention strategies. 2. Materials 2.1. Immunoafﬁnity Depletion of High-Abundant Proteins 2.1.1. Serum Collection 1. Becton Dickinson Vacutainer tubes (VWR International) with SST gel and BD clot activator.

2.1.2. Immunoafﬁnity Chromatography 1. An 1100 liquid chromatograph (Agilent Technologies, Wilmington, DE.) consisting of a binary gradient pumping system, a thermostatted autosampler,

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a variable wavelength absorbance detector, a temperature controlled column compartment, and a thermostatted automated analytical scale fraction collector (see Note 1). 2. Agilent Chemstation Software version B.01.03. 3. Multiple Affinity Removal System column, 4.6 mm id × 100 mm, Immunodepletion Buffer A and Buffer B (Agilent Technologies, Wilmington, DE). 4. 0.22-μm spin filters.

2.2. Immunodepleted Serum and Bound Fraction Processing 1. BCA Assay Kit (Pierce, Rockford, IL). 2. Tris-glycine precast gels (4–20% acrylamide, 10 wells, 1 mm), sample preparation (loading) and running buffers, Mark12 unstained standards (Invitrogen, Carlsbad, CA). 3. 4-mL spin concentrators with 5 kDa molecular weight cutoffs (Agilent Technologies, Wilmington, DE). 4. Standard equipment for two-dimensional electrophoresis (Bio-Rad, Hercules, CA). 5. Rehydration buffer: 8 M urea, 2% CHAPS, 2% ampholytes and 20 mM dithiothreitol. 6. 11 cm immobilized pH gradient (IPG), pH 3–10 nonlinear strips (Bio-Rad, Hercules, CA). 7. 8–16% precast Tris-glycine gels for second dimension separation (Bio-Rad). 8. Gel Code Blue – Coomassie stain (Pierce, Rockford, IL).

2.3. Reversed-phase Separation of Low-abundant Proteins 1. An 1100 liquid chromatograph (Agilent Technologies, Wilmington, DE.) consisting of a binary gradient pumping system equipped with a 900 μL capillary injector loop, a thermostated autosampler, a variable wavelength absorbance detector, a temperature controlled column compartment, and a thermostated automated analytical scale fraction collector (see Note 1). 2. 4.6 mm ID × 50 mm macroporous high recovery reversed-phase C18 column (mRPC18) (Agilent Technologies, Wilmington, DE). Particle composition is a silica-based macroparticulate material with a particle size of 5.0 μm. The column hardware is made of PEEK composition and the frits are 2.0 μm PEEK encapsulated.

3. Methods 3.1. Immunoafﬁnity Depletion of High-Abundant Proteins 3.1.1. Serum Collection 1. Collect serum samples into a Becton Dickinson Vacutainer tubes (VWR International) with SST gel and BD clot activator. After clot formation, centrifuge sample at 1000g for 15 min. Remove serum and store aliquotes at –80°C. Total time for serum processing is less than 60 min.

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31

3.1.2. Immunoafﬁnity Chromatography The immunoaffinity column depletion technology offers rapid and simultaneous removal of six high-abundant proteins in 28 min per sample (Fig. 3). The column is based on the rabbit polyclonal antibodies to six major serum proteins—human serum albumin (HSA), transferrin, alpha1-anti-trypsin, haptoglobin, immunoglobulin A (IgA), and immunoglobulin G (IgG) that were affinity purified on corresponding protein antigen columns (4). The resulting affinity-purified antibodies were covalently coupled to porous beads via their Fc region and cross-linked. Spatially controlled cross-linking of the antibodies resulted in preferential orientation of the antibody binding sites away from the solid-phase surface, supporting maximum binding capacity of targeted proteins. Through the series of column loading, washing, collection, and reequilibration steps, multiple serum samples can be processed and depleted of targeted high-abundant proteins. After each pass of serum through the column, flow-through fractions containing low-abundant proteins can be pooled and concentrated for downstream postaffinity processing. The methods outlined below demonstrate a typical workflow for immunodepleting a given quantity of human serum samples. Similar procedures have been employed successfully for cerebrospinal fluid, amniotic fluid and for urine analysis. This methodology is robust, scalable, and easily automated for multiple samples processing, as well as compatible with downstream 1D and 2D-SDSPAGE, LC, LC/MS, and/or enzymatic or chemical fragmentation methods. 1. Purge LC system with Buffer A and Buffer B at a flow rate of 1.0 mL/min for 10 min. without column (see Note 2).

Bound Fraction

Serum Injection

Flow-through Fraction Elution

Re-equilibration

Fig. 3. Chromatogram for the affinity removal of high-abundant proteins from human serum. 35 μL of serum was diluted 5× and injected on a 4.6 mm id × 100 mm immunoaffinity column (0.50 mL/min) and a flow-through peak (3–5.0 min) was collected for reversed-phase HPLC fractionation. The column was washed with Buffer A and the targeted high-abundant proteins were eluted with Buffer B.

32

Martosella and Zolotarjova Table 1 LC timetable

1 2 3 4 5 6

Time (mins)

%B

Flow rate

0.00 10.00 10.01 17.00 17.01 28.00

000 000 10000 10000 000 000

0.500 0.500 1.000 1.000 1.000 1.000

Max. pressure 120 120 120 120 120 120

2. Run two method blanks by injecting 200 μL of Buffer A without the column according to the LC timetable (Table 1) (see Note 3). Table 1 here 3. Attach a 4.6 × 100 mm immunoaffinity column and equilibrate in Buffer A for 4 min at a flow rate of 1 mL/min. 4. Dilute human serum five times with Buffer A (for example 35 μL of human serum with 140 μL of Buffer A (see Note 4). 5. Remove sample particulates with a 0.22 μm spin filter and centrifuge sample for 1.0 min at 16,000g. 6. Inject 175 μL of the diluted serum at a flow rate of 0.5 mL/min and run LC timetable. 7. Collect the flow-through fraction (appears between 3.0–5.0 min; see Fig. 3 for the chromatogram) into 1.5-mL microcentrifuge tubes at 4°C. Store collected fraction at –20°C if not analyzed immediately. 8. Wash column with buffer A (see LC timetable) and elute the bound fraction with Buffer B at a flow rate of 1.0 mL/min for 7.0 min. 9. Regenerate the column by equilibrating it with Buffer A for 11.0 min at a flow rate of 1.0 mL/min for a total run cycle of 28.0 min. (see Note 5).

3.2. Immunodepleted Serum and Bound Fraction Processing For analyses of immunodepleted serum before RP separation or without further RP fractionation proceed with the recommendations below. For direct processing of the flow-through for RP HPLC fractionation of the immunodepleted serum see Section 3.3. 1. If lyophylization of the immunodepleted serum is desired, buffer exchange to a volatile buffer (for example, ammonium bicarbonate) because of high salt concentration in Buffer A. 2. Measure protein concentration in serum, flow-through and bound fractions using BCA protein assay. For 1D-SDS-PAGE analysis of the flow-through fraction (immunodepleted serum) or bound fraction, mix sample aliquots (5–10 μg of protein) with the equal volume of the loading buffer, boil sample for 3 min. and

Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase 1

2

3

4

33

5

200.0 116.3 97.4 66.3 55.4 36.5 31.0 21.5 14.4 6.0 3.5

Fig. 4. 1D SDS gel electrophoresis of human serum protein fractions from an immunoaffinity column. An equal amount (9 μg) of crude serum (Lane 2), flow-through (Lane 3) and bound fractions (Lane 4) were separated on 4–20% SDS-PAGE gel under nonreducing conditions. Lanes 1 and 5 are the molecular weight standards (Mark12) from Invitrogen. The proteins were stained with Coomassie Blue dye. Based on the protein assay of the flow-through fraction, 85% of total protein was removed from the crude serum.

load on the gel. Fig. 4 shows 1D gel electrophoresis data for crude serum, flowthrough and bound fractions from an immunoaffinity column. Equal amounts of protein (9 μg) were loaded in each lane. Results show that high-abundant proteins in serum (Lane 2) are clearly removed and are not visible in the flowthrough fraction (Lane 3). Also, low-abundant proteins that were not visible in the serum before depletion (Lane 2) became visible in the flow-through fraction after removal of the high-abundant proteins. 3. For IEF, 2D-SDS-PAGE, and MS analysis, it is necessary to buffer exchange/desalt and concentrate fractions to an appropriate buffer. Use 4 mL spin concentrators with 5 kDa molecular weight cutoffs. Centrifuge samples at 7,500g for 20 min at 4°C. Buffer exchange into 20 mM Tris-HCl, pH 7.4, by 3 rounds of buffer addition, with 20 min centrifugation for each round. Aliquot the concentrated samples and store at –70°C until the analysis. Analyze protein concentration using a BCA protein assay. 4. Prepare 2D electrophoresis samples by mixing 250 μg of proteins with 185 μL of rehydration buffer containing 8 M urea, 2% CHAPS, 2% ampholytes, pH 3–10, and 20 mM dithiothreitol. Apply samples on 11-cm immobilized pH gradient (IPG), pH 3–10 nonlinear strips and process them according to

34

Martosella and Zolotarjova the manufacturer’s instructions. Perform the second dimension separation on 8–16% precast Tris-glycine gels. Visualize proteins by Coomassie Blue staining. Fig. 5 shows the protein pattern of human serum before (Panel A) and after immunodepletion (Panel B). Circles indicate the areas where the targeted highabundant proteins reside. The depletion of high-abundant proteins unmasks the low-abundant proteins because of the substantial removal of protein mass from the sample. More than 85% of total protein was depleted after a single pass of serum through the immunoaffinity column. This enabled a large increase in low-abundant protein mass loading onto the gel (up to 10 times). As a result, lowabundant protein fractions become enriched and more easily detectable on the gel, making the protein spots more amenable to quantitation and MS identification.

The immunoaffinity column is highly specific for the removal of highabundant proteins from serum (5). A small number of nontargeted proteins are bound to the column. None of the nonspecific proteins are bound quantitatively to the immunoaffinity column and represent only a small percentage of the total flow-through.

A, Crude Serum

B, Serum after immunodepletion

Anti-trypsin IgA Transferrin IgG Heavy Albumin MW Chain

(kDa) 200 116 97 66 55

Haptoglobin

MW (kDa) 200 116 97 66 55

Ig Light 37 Chain 31

37 31

21

21

14

14

6

6

pH 3 –10

Fig. 5. 2D gel electrophoresis of human serum before and after removal of highabundant proteins. Panel A. Human serum before depletion. The targeted high abundant proteins are circled. Panel B. Human serum after depletion of the six targeted high abundant proteins. The positions of the removed proteins are circled. 250 μg of total protein loaded on each gel. Molecular weight standards–Mark12 (Invitrogen). Proteins were visualized by staining with Coomassie Blue.

Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase

35

3.3. Reversed-phase Separation of Low-Abundant Proteins 3.3.1. Reversed-Phase Chromatographic Conditions 1. Eluent A: 0.1% TFA in water, Eluent B: 0.08% TFA in acetonitrile. 2. Autosampler and fraction collection temperature: 4°C. 3. Column temperature: 80°C. If column oven cannot reach or maintain 80°C, a lower temperature can be used, however, protein recovery and chromatographic resolution may be increasingly compromised with decreasing temperature. 4. UV absorbance: 280 nm (preferred) or 210 nm. 5. Sample flow rate: 0.75 mL/min. 6. Prepare LC gradient elution according to LC Timetable in Table 2. 7. Prepare flow-through sample directly for Reversed-Phase chromatographic separation (see Note 6). Allow sample to equilibrate at room temperature for at least 30 min. 8. Inject up to 900 μL of the denatured (6M urea/1.0% acetic acid) flow-through proteins from immunodepletion. If it is desirable to process the entire flowthrough sample volume from each immunodepletion run (for example 1.5 mL of denaturated flow-through) or load multiple flow-throughs, an injector loading program is needed. This can also be useful for utilizing the columns full loading capacity when flow-throughs from multiple immunodepletions have been pooled (see Note 7). Fig. 6 is representative of a RP elution profile of immunodepleted human serum obtained when using the optimized multi-segmented elution conditions presented in Table 2. The RP separation can be fractionated and the collected fractions processed for downstream electrophoretic or LC/MS analyses (6). 9. Collect fractions at 1.0 min time intervals from 0–54 min. At the recommended flow of 0.75 mL/min., the fraction volumes will be 0.75 mL (see Note 8). 10. Dry each fraction in a centrifugal vacuum concentrator. To avoid protein degradation do not dry overnight. If sample processing is not immediate, store dried fractions at –80°C. 11. For SDS-PAGE analysis of the separation efficiency (an example of the results produced is shown in Fig. 7), dry fractions in the vacuum concentrator and Table 2 LC timetable

1 2 3 4 5 6 7 postrun re-equilibration

Time (mins)

%B

0.00 6.00 39.0 49.0 53.0 58.00 68.00

3.00 30.0 55.0 100.0 100.0 3.00 3.00

36

Martosella and Zolotarjova

mAU

Absorbance @ 280 nm (mAu)

35 30 25 20 15 10 5 0 0

5

10

15

20

25 30 Time (min.)

35

40

45

min

Fig. 6. Representative RP-HPLC elution profile (absorbance at 280 nm) for human serum depleted of high-abundant proteins. An aqueous TFA and ACN (TFA) gradient was used at 80°C at a flowrate of 0.75 mL/min on a 4.6 mm id × 50mm mRP-C18 column. The sample comprised a total of 270 μg protein in 6 M urea/1% AcOH. then dissolve each with electrophoresis sample preparation (loading) buffer as recommended below (see Note 9). 12. For 2D PAGE analysis of the RP fractions, we suggest combining the fractions in a manner suitable to accommodate specific workflows and goals. Fraction pooling will be required to achieve appropriate protein loads for 2D PAGE analysis. To process the RP fractions, dry in a vacuum concentrator and dilute with the IEF buffer according to the manufacturers protocol. 13. For consistent and repeatable reversed-phase separation consult Note 10.

4. Notes 1. Conventional autosamplers generally do not provide optimum sampling because of conditions which can lead to extra column band broadening and mixing, thereby reducing resolution. 2. The immunoaffinity column requires a proprietary two buffer system (Buffer A and Buffer B) for operation. The two buffers provide the means to separate

Multi-Component Immunoafﬁnity Subtraction and Reversed-Phase Fractions

7

8

9

10 11

12 13 14 15 16

17 18 19 20 21 22 23 24 25

37 26

kDa 200.0 116.3 97.4 66.3 55.4 36.5 31.0 21.5 14.4

Fractions

27 28 29 30 31 32 33 34 35 36

kDa 200.0 116.3 97.4 66.3 55.4 36.5 31.0 21.5 14.4

Fig. 7. SDS-PAGE analysis of RP-HPLC fractionated immunodepleted human serum from an mRP-C18 column (4.6 mm id × 50 mm). A depleted human serum sample was injected onto the column and eluted by a multi-segment gradient. Fifty-four fractions were collected (29 showing the majority of protein elution) for analysis by 4–20% SDS-PAGE.

low-abundant proteins from high-abundant proteins and regenerate the column, all in about 20–30 min per injection. Buffers A and B are optimized to minimize co-adsorption of nontargeted proteins to the column packing, and to ensure reproducibility of column performance and long column lifetime. Buffer A is a salt-containing neutral buffer (pH 7.4) used for loading, washing, and reequilibrating the column. Buffer B is a low pH urea buffer used for eluting the bound high-abundant proteins from the column. Serum samples are injected onto the column and the high-abundant proteins are simultaneously removed as low-abundant proteins pass through in the flow-through fraction. After collecting the low-abundant proteins and washing the column, the bound proteins are eluted with Buffer B and the column is re-equilibrated with Buffer A. Do not expose immunoaffinity column to solvents other than Buffers A and B. Organic solvents (acetonitrile, alcohols, etc.), strong oxidizers, acids, reducing agents, and other protein denaturing agents will cause irreversible column damage.

38

Martosella and Zolotarjova

3. 4.

5. 6.

7.

8.

Under optimized operating conditions the column stable and robust for at least 200 injections of serum samples. Ensure proper sample loop size in autosampler. Consult column certificate of analysis to verify column capacity. Concentrations of some high-abundant proteins can vary widely depending on the sample source. Proteins such as alpha1-antitrypsin, haptoglobin, IgG rise several folds in response to stress, infection, inflammation, or tissue necrosis and are known as acute phase reactants. Users need to adjust column loading volume accordingly. It is not recommended to load crude serum onto the column. Addition of protease inhibitors cocktail (Complete, Roche, IN) in buffer A for sample dilution helps prevent protein degradation. When not in use, store the column, with the end-caps sealed, in a refrigerator at 2–8°C to minimize losses in column capacity. Do not freeze the column. Flow-through sample preparation: The amount of flow-through sample volume from a 4.6 mm id × 100 mm immunodepletion column is approx 1.0–1.5 mL. Immunodepleted serum fractions can either be pooled together or processed individually. However, by either method, they must first be denatured under acidic conditions. To denature before RP separation, 480 mg solid urea and 13 μL acetic acid (AcOH) are added for every 1.0 mL of flow-through for a final sample concentration of approx 6 M urea/1.0% AcOH. Calculation is based on an immunodepletion separation of 35 μL human serum (diluted 5×), in which BCA protein analysis gave a flow-through protein concentration of 0.38 mg/mL. It is recommended to measure the actual flow-through protein concentration for each serum sample lot processed and adjust the 6 M urea and 1.0% AcOH concentrations accordingly. For loading sample volumes greater than 900 μL, an isocratic loading method is required. To load under isocratic conditions set 97% Eluent A (0.1% TFA in water) and Eluent B (0.08% TFA in acetonitrile) at 3.0% for a minimum run time of 3.0 min (maintain all other chromatographic conditions from Section 3.3). Perform desired amount of column injections without overloading the column. The maximum loading capacity for a 4.6 mm id × 50 mm mRP-C18 column is approx 400 μg of immunodepleted serum. After multiple sample injections have been loaded, proceed with the elution gradient in Table 2. In Chemstation, this process can be automated and configured under Sequence and Sequence Table. Depending on the user’s capacity for sampling handling and/or processing objectives, some workflows may require the need to collect more or a lesser amount of fractions and may therefore need to vary the time-based collection. However, fraction collection greater than 2.0 min time intervals will require collection tubes larger than 2.0 mL and typically require tray or instrument adjustments for many automated fraction collectors. Fractions collected from 1 to 6 min and 37 to 54 min are not shown in Fig. 7. The majority of protein elution as determined by Commassie blue staining occurs from 7 to 36 min. If a comprehensive MS analysis is the goal, we recommend fraction collection throughout the entire run.

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39

9. For fractions #1–#12 and #37–#54, dissolve each fraction in 30 μL of electrophoresis sample preparation buffer and heat for 3 min at 70°C. If the entire protein load on the column was 400 μg or less, load the entire 30 μL of prepared sample directly onto the gel. If protein fractions were pooled from several chromatographic runs and thus exceeded 400 μg of total column load, remove 15 μL of prepared sample and dilute with 15 μL deionized water (30 μL total) and load onto the 1D SDS-PAGE. For fractions #13–#36, dissolve each fraction in 75 μL of sample preparation buffer and heat for 3 min at 70°C. If the entire protein load on the column was 400 μg or less, directly load 15 μL (without water dilution) onto SDS-PAGE. If protein fractions were pooled from several chromatographic runs and thus exceeded 400 μg of total column load, remove 7 μl of prepared sample, dilute with 7 μL deionized water (14 μL total) and load onto the gel. 10. Column runs from the same depleted serum sample, during the same run progression, should be very repeatable with identical peak shapes and intensities. If variances are occurring the column may need replacing. Column life varies with use and conditions, but should typically last for over 75 injections. It is recommended to complete each RP separation with a minimum postrun time of 10.0 min to ensure that the column has re-equilibrated. Periodically perform blank injections to evaluate baseline stabilization. If peak ghosting, which is a characteristic of protein carryover, is present, perform a run with 100% Eluent B for 4 min, maintain elevated temperature, and repeat the blank injection. If ghosting is still present, yet minimized, repeat the run of 100% Eluent B. When not in use store column at room temperature in 25–75% (v/v) water-methanol with the column ends capped.

References 1. Anderson, N. L., Anderson, N. G., (2002) The human plasma proteome: history, character, and diagnostic prospects. Mol Cell Proteomics, 1, 845–67. 2. Duan, X., Yarmush, D. M., Berthiaume, F., Jayaraman, A., and Yarmush, M. L. (2004) A moose serum two-dimensional gel map: application to profiling boon injury and infection. Electrophoresis, 25, 3055–65. 3. Fujii, K., Nakano, T., Kawamura, T., Usui, F., Bando, Y., Wang, R., and Nishimura, T. (2004) Multidimensional protein profiling technology and its application to human plasma proteome. J.Proteome Res., 3, 712–18. 4. Zolotarjova, N., Martosella, J., Nicol, G., Bailey, J., Boyes, B.E., and Barrett, W.C. (2005) Differences among techniques for high-abundant protein depletion. Proteomics, 5, 3304–13. 5. Harlow, E. and Lane, D., (1988) Antibodies, A Laboratory Manual. Cold Springs Harbor Laboratory: New York, pp. 726. 6. Martosella, J., Zolotarjova, N., Liu, H., Nicol, G., and Boyes, B.E., (2005) Reversedphase high-performance liquid chromatographic prefractionation of immunodepleted human serum proteins to enhance mass spectrometry: identification of lowerabundant proteins. J. Proteome Res. 4, 1522–1537.

4 Immunoafﬁnity Fractionation of Plasma Proteins by Chicken IgY Antibodies Lei Huang and Xiangming Fang

Summary Separation of complex mixtures having a wide dynamic range of protein concentration, such as plasma or serum, presents a significant challenge for proteomic analysis. Immunoaffinity fractionation is one of the most effective methods used during sample preparation to improve the ability to detect low-abundant proteins (LAP), enhancing biomarker discovery. Avian IgY (Immunoglobulin Yolk) antibodies have unique and advantageous features, which include strong avidity, high specificity, low nonspecific binding, and accumulative production. Polyclonal IgY antibodies covalently coupled to microbeads are particularly effective in specifically removing high-abundant proteins (HAP) from plasma, serum, CSF, urine, and other body fluid or cellular sources. IgY-12 is a composition of IgY microbeads designed for one-step removal of the 12 most abundant proteins in human serum or plasma: albumin, IgG, transferrin, fibrinogen, 1antitrypsin, IgA, IgM, 2-macroglobulin, haptoglobin, apolipoproteins A-I and A-II, and orosomucoid (1-acid glycoprotein). Removal of the 12 HAPs enables improved resolution and dynamic range for one-dimensional gel electrophoresis (1DGE), two-dimensional gel electrophoresis (2DGE), and liquid chromatography/mass spectrometry (LC/MS).

Key Words: High abundance proteins; IgY antibodies; immunoaffinity fractionation; low abundant proteins; protein depletion; protein separation; proteomics; sample preparation.

1. Introduction IgY antibody is immunoglobulin gamma isolated from egg yolks (so called IgY) of certain avian and reptilian vertebrates such as birds, reptiles, and amphibians (1–3). Chicken IgY antibodies have been developed and From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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successfully applied for various types of immunoassays (4–8). An outstanding advantage of chicken IgY antibodies is that they are secreted by hens into egg yolk, resulting in a high-yielding and easy to access reservoir of antibodies (9). Compared to drawing blood, collecting eggs is noninvasive, continuous, convenient, and scalable. One egg contains about 100 mg of total IgY. A laying hen can actively produce eggs for 2 years at an average of 20 eggs per month. Distinct from mammalian IgG antibodies in molecular structure and biochemical features, IgY antibodies have been shown to have several advantages over IgG, particularly for their high avidity and less cross-reactivity to human proteins (10–12). This is because of the avian affinity maturation mechanism of gene conversion and the great evolutionary distance between chicken and mammals. When mammalian protein antigens are used to immunize chickens, more immunogenic epitopes are presented to the host, resulting in IgY antibodies with high affinity and broader recognition spectrum. In addition, the IgY Fc region does not bind human proteins such as complement, rheumatoid factor, Fc receptor, etc, thus significantly increasing IgY’s specificity of capture. Another unique feature of IgY antibodies is that they have a broader antigenbinding host range. This is also the result of greater evolutionary distance between chickens and mammals, and the sequence similarity among mammals. IgY antibodies raised against these high abundance proteins using human antigens also recognize the orthologous proteins from other mammalian species such as nonhuman primates, rat, mouse, pig, goat, cow, and dog. IgY microbeads are produced by covalently coupling IgY antibodies to 60-μm polymeric beads via the oligosaccharides located on their Fc region. This orientated conjugation allows maximal capture of target proteins. Compared to other affinity reagents, including IgG microbead products, IgY microbeads have been shown to have distinct features and advantages (13,14). IgY-12 is a mixture of 12 types of IgY microbeads designed to collectively remove albumin, IgG, 1-antitrypsin, IgA, IgM, transferrin, haptoglobin, 1-acid glycoprotein (orosomucoid), 2-Macroglobulin, HDL (mainly apolipoproteins A-I and AII), and fibrinogen from complex human body fluids such as serum, plasma, and cerebral spinal fluid (CSF) in a single step. IgY-12 spin column (0.6-mL bed size) can process 15–20 μL human plasma per loading, yielding 100–160 μg of proteins partitioned of HAP. Larger samples can be processed by IgY12 liquid chromatography (LC) columns. A 2-mL LC column can partition 40–50 μL human plasma per injection and a 10-mL LC column allows a single loading of 200–250 μL. Through a simple procedure of sample loading, washing, eluting, and regenerating, approx 90–95% of total protein mass from human serum or plasma is removed. The LAP in the flow-through fractions can be further studied by 2D polyacrylamide gel electrophoresis (PAGE) or LC/MS. The regenerated beads can be reused many times with minimal protein

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carry-over (15). The IgY microbeads can also be used in 96-well filter plate and other formats for high-throughput partitioning of human serum/plasma samples or other body fluids.

2. Materials 2.1. IgY-12 High Capacity Spin Column Kit 1. Prepacked IgY-12 spin column, containing 1.2-mL IgY microbeads slurry (Beckman Coulter, Fullerton, CA). Store at 2–8°C. Do not freeze. 2. Dilution buffer (Tris Buffered Saline, TBS): 10 mM Tris-HCl, pH 7.4, 150 mM NaCl. For sample dilution, washing and equilibrating column, and rinsing pipet tips during resin transfer. Store at room temperature. 3. Stripping buffer: 0.1M Glycine-HCl, pH 2.5. For stripping off bound proteins from column. Store at room temperature. 4. Neutralization buffer: 1M Tris-HCl, pH 8.0. For neutralizing column and eluted proteins. Store at room temperature. 5. 2-mL collection tubes. For collecting flow-through, washing, and eluted fractions. 6. Empty spin columns with end caps.

2.2. IgY-12 High Capacity LC2 or LC10 Column Kit 1. Prepacked IgY-12 LC column, 2-mL or 10-mL packed bed (Beckman Coulter, Fullerton, CA). Store at 2–8°C. Do not freeze. 2. Dilution, stripping, and neutralization buffers are same as for spin column. 3. Spin filters. For sample clean up before loading column to remove sample particulates and extend column life.

2.3. IgY-12 Microbeads for 96-Well Filter Plates 1. IgY-12 microbeads, 50% slurry (GenWay Biotech, San Diego, CA). Store at 2–8°C. Do not freeze. 2. Dilution, stripping, and neutralization buffers are same as for spin column. 3. 96-well filter plate, 400 μL, UHMW PE 25 μM, Long drip. (Innovative Microplate, Billerica, MA). Store at room temperature. 4. Collection plate, Nunc 96-DeepWell™ Plates, 1.2-mL, Polypropylene (NUNC, Rochester, NY).

2.4. SDS Gel Electrophoresis 1. Tris-HCl SDS Gel, precast, 4–20% linear gradient (Rio-Rad, Hercules, CA). Store at 2–8°C. 2. 5× SDS Sample Buffer: 10% (w/v) SDS, 20 mM dithiothreitol (DTT) or 25% (w/v) -mercaptoethanol (BME) (omitted under nonreducing condition), 20% (w/v)

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glycerol, 0.2M Tris-HCl, pH 6.8, 0.05% (w/v) bromophenol blue. Store at room temperature. 3. Tris/glycine/SDS electrophoresis buffer: 25 mM Tris-base, 200 mM glycine, 0.1% (w/v) SDS. 4. Coomassie Blue Staining Solution: 0.25% (w/v) Coomassie Brilliant Blue R250, 40% (v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 5. Destain Soluton: 40% (v/v) methanol, 10% acetic acid. Store at room temperature.

3. Methods 3.1. Spin Column (see Note 1) 3.1.1. Immunocapture of 12 Abundant Serum/Plasma Proteins 1. Dilute 15–20 μL serum or plasma sample in dilution buffer to obtain a final volume of 600 μL. 2. Snap off the tip from the column and place the column in a 2-mL collection tube. 3. Centrifuge the column for 30 sec at 400g in a microcentrifuge to obtain dried beads. 4. Place the end cap to the column. Immediately add 0.5 mL diluted sample to the dried beads in the column. Seal the column with the top snap cap. 5. Mix the beads and the sample completely by inverting and shaking the column, place it on an end-to-end rotator and incubate at room temperature for 15 min. 6. Invert the column. Remove the end cap and place the column in a 2-mL collection tube. Centrifuge for 30 sec at 400g. Collect flow-through (IgY-12-depleted) sample for further analysis (see Note 2).

3.1.2. Washing of Column 1. Wash beads with 0.5 mL of dilution buffer, a total of three times. To obtain maximum yields of flow-through samples, the fraction from the first washing can be collected and combine with the flow-through sample from Section 3.1.1 step 6 for further analysis. 2. For each wash, always first insert the end cap, and then add 0.5 mL of dilution buffer and seal the column with top snap cap. Mix the beads and buffer completely by inverting and shaking the column, remove the end cap while inverting the column and place it in a 2-mL collection tube. Centrifuge for 30 sec at 400g and save the flow-through for further analysis.

3.1.3. Stripping of Bound Proteins 1. Strip off bound proteins from beads using stripping buffer, a total of three to four times. For each elution, place the end cap to the column first after centrifugation, then add 0.5-mL stripping buffer and seal the column with top snap cap. Mix the beads and buffer completely (see Note 3) by inverting and shaking the column, incubate at room temperature for 3 min, remove the end cap while holding the

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column upside down and place it in a 2-mL collection tubes. Centrifuge for 30 sec at 400g and collect the eluate. It is crucial for column stability to immediately neutralize the beads (see Section 3.1.4.). 2. Pool eluted samples (total 1.5–2.0 mL) and neutralize with 150–200 μL of neutralization buffer. Samples can be stored at -80°C if not analyzed immediately.

3.1.4. Regeneration of IgY-12 Microbeads 1. To regenerate IgY-12 microbeads after stripping bound serum proteins as described above, immediately neutralize beads with 0.6 mL of 1:10 diluted neutralization buffer. Mix beads and buffer completely by inverting and shaking the column. Incubate at room temperature for 5 min. Spin down beads in the column for 30 sec at 400g. 2. Resuspend beads in 0.5-mL of dilution buffer. Beads are ready for next cycle or storage at 4°C. For storage of regenerated beads, add sodium azide (NaN3 ) to 0.02%. (w/v) in dilution buffer.

3.2. LC Column (see Note 4) 3.2.1. Protocol for 6.4 × 63 mm (2 mL) Column 1. Set up the three Buffers as the only mobile phases. 2. Purge lines with three Buffers at a flow rate of 1.0 mL/min for 10 min. 3. Set up LC timetable (see Table 1 for details) and run two method blanks by injecting 125 μL of dilution buffer without a column. 4. Attach column and equilibrate it in dilution buffer for 10 min at a flow rate of 1.0 mL/min at room temperature. 5. Dilute human serum five times (for example: 50 μL human serum with 200 μL of dilution buffer). 6. Remove particulates with a 0.45-μm spin filter; 1 min at 9,000g. 7. Inject 250 μL of the diluted and filtered plasma sample (Column capacity: 40–50 μL of neat human serum/plasma per injection), start the method at a flow rate of 0.1 mL/min for 10 min, wash the column at a flow rate of 0.2 mL/min for 7 min, then change the flow rate to 1.0 mL/min to continue the wash for 5 min, collect flow-through fraction and store collected fractions at –80°C if not analyzed immediately. 8. Elute bound proteins from the column with stripping buffer at a flow rate of 1.0 mL/min for 142 min, and neutralize the column with neutralizing buffer at a flow rate of 1.0 mL/min for 6 min. 9. Regenerate column by equilibrating it with dilution buffer for an additional 6 min at a flow rate of 1.0 mL/min. 10. Store column after equilibrating with dilution buffer containing 0.02% (w/v) sodium azide (NaN3 ) at 2–8°C in a refrigerator. 11. A standard chromatograph is illustrated in Fig. 1.

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Table 1 LC Method for a 6.4 × 63 mm column Cycle

Injection Wash Wash Wash Elution Neutralization Re-equilibrium Stop

Time (min)

0 10.01 17.01 22.01 36.01 42.01 48.00

Dilution Stripping Neutralization Flow rate buffer buffer buffer (mL/min)

100 100 100 0 0 100

0 0 0 100 0 0

0 0 0 0 100 0

0.1 0.2 1.0 1.0 1.0 1.0

Max pressure (psi) 100 100 100 100 100 100

Optimized for Beckman System Gold HPLC, Pump Module 1 Type: 118, Detector Model: 166

3.2.2. Protocol for 12.7 × 79.0 mm (10 mL) Column 1. Set up the three buffers as the only mobile phases. 2. Purge lines with three buffers at a flow rate of 1.0 mL/min for 10 min. 3. Set up LC timetable (see Table 2 for details) and run two method blanks by injecting 1.25 mL of dilution buffer without a column.

Fig. 1. Chromatography of immunoaffinity separation of human plasma using IgY12 high capacity LC2 column. Fifty microliters human plasma was fractionated on the column.

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4. Attach column and equilibrate it in dilution buffer for 10 min at a flow rate of 2.0 mL/min at room temperature. 5. Dilute human serum/plasma five times (for example: 250 μL human plasma with 1.0 mL of dilution buffer). 6. Remove particulates with a 0.45 μm spin filter; 1 min at 9,000g. 7. Inject 1.5 mL of the diluted and filtered plasma sample (column capacity: 200–250 μL of neat human serum/plasma), start the method at a flow rate of 0.5 mL/min for 30 min, wash the column at a flow rate of 2.0 mL/min for 5 min, collect flow-through fraction and store collected fractions at -80°C if not analyzed immediately. 8. Elute bound proteins from the column with stripping buffer at a flow rate of 2.0 mL/min for 15 min, and neutralize the column with neutralizing buffer at a flow rate of 2.0 mL/min for 10 min. 9. Regenerate column by equilibrating it with dilution buffer for an additional 10 min at a flow rate of 2.0 mL/min. 10. Store column after equilibrating with dilution buffer containing 0.02% (w/v) sodium azide (NaN3 ) at 2–8°C in a refrigerator. 11. A standard chromatograph is illustrated in Fig. 2

3.3. 96-Well Spin Filter Plate 1. For each well, dilute 2–3 μL human serum or plasma in dilution buffer to obtain final volume of 100 μL. 2. Aliquot 200 μL per well IgY-12 microbeads slurry into 96-well filter plate. Spin plate at 190g for 1 min in Eppendorf bench top centrifuge (see Note 5) with plate adapter to remove buffer. 3. Add diluted sample to each well, mix with pipet tip. Incubate at room temperature on shaker for 15 min. Table 2 LC method for a 12.7 × 79.0 mm column Cycle

Injection Wash Wash Elution Neutralization Re-equilibrium Stop

Time (min)

0 30.01 35.01 50.01 60.01 70.00

Dilution Stripping Neutralization Flow rate buffer buffer buffer (mL/min)

100 100 0 0 100

0 0 100 0 0

0 0 0 100 0

0.5 2.0 2.0 2.0 2.0

Max pressure (psi) 100 100 100 100 100

Optimized for Beckman System Gold HPLC, Pump Module 1 Type: 118, Detector Model: 166

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Fig. 2. Chromatography of immunoaffinity separation of human plasma using IgY12 LC10 column. Two hundred fifty microliters human plasma was fractionated on the column. 4. Spin plate at 190g for 1 min. Collect flow-through fraction in collection plate, about 100 μL. 5. Add 100 μL dilution buffer to each well. Gently shake the plate and spin at 190g for 1 min. Collect flow-through faction into the same collection plate from step 4. 6. Wash beads with 100 μL dilution buffer, a total of 3 times. For each wash, add 100 μL dilution buffer to each well, gently shake the plate and spin at 190g for 1 min. Collect flow-through faction into collection plate for future analysis. 7. Add 100 μL stripping buffer to each well. Gently shake the plate and incubate at room temperature on shaker for 2 min. Spin the plate at 190g for 1 min. Repeat for three to four times. Collect and combine flow-through factions into collection plate for future analysis. 8. Immediately add 100 μL neutralization buffer to each well. Gently shake the plate and incubate at room temperature on shaker for 5 min. 9. Spin the plate at 190g for 1 min. Add 100 μL dilution buffer to each well. Beads are ready for next cycle or storage at 4°C. For storage of regenerated beads, add sodium azide (NaN3 ) to 0.02%. (w/v) in dilution buffer.

3.4. Evaluation of Fractionation Efﬁciency by SDS-PAGE (see Note 6) 1. Take a small aliquot of samples from neat plasma, flow-through and eluted fraction. Mix with 5× SDS sample buffer. Load approx 25–30 μL (see Note 7)

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Fig. 3. SDS-PAGE analysis of neat plasma, flow-through and eluted fractions using IgY-12 high capacity spin column (1.0 mL slurry). Twenty microliters of human plasma was partitioned on the column. Five cycles were repeated. Fifteen microliters of 1:70 dilution of neat plasma, 15 μL of flow-through fraction, and 15 μL of eluted fraction were loaded to 4–20% SDS gradient gel. Coomassie blue staining. M: molecular weight marker, S: Neat plasma. F1-F5: Flow-through fractions from cycle 1 to 5, E1-E5: Eluted/bound fractions from cycle 1 to 5.

of each sample on 4–20% SDS gel. The gel is run in Tris/Glycine/SDS electrophoresis buffer at 200 volts for 35 min. 2. Remove gel from the gel cassette. Rinse the gel with deionized water. Stain gel in Coomassie Blue Staining solution for 30 min to 1 h on shaker. 3. Rinse the gel with deionized water. Place gel in destaining solution and shake until the bands emerge clearly from the background. Replace destaining solution with deionized water. 4. A successful fractionation will result in distinct banding patterns on the gel as shown in Fig. 3. The major protein bands of albumin, transferrin, IgG, and Apo-A1 that disappeared in the flow-through fraction will be shown in the eluted fraction.

4. Notes 1. Before loading plasma sample to the new IgY-12 Spin Column, perform the full procedure with buffers only for one or two cycles. The purpose is to remove any residual uncoupled IgY antibodies in the column. 2. The flow-through fraction is now greatly diluted. The samples can be concentrated to desired concentration and volume for downstream analysis using molecular

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4.

5. 6.

7.

8.

9.

Huang and Fang weight cutoff centrifugal concentrators, such as Vivaspin (Sartorius, Goettingen, Germany). An alternative method is TCA/Acetone precipitation. The beads tend to be packed tighter in acidic solution after centrifugation. Make sure beads are resuspended well for effective stripping. For more efficient stripping, 0.25 M Glycine-HCl, pH 2.5 can be used. In this case, 1 to 4 dilution of neutralization buffer should be used to regenerate beads and 250 μL of neutralization buffer should be added per 1 mL of eluted samples. The LC protocols are optimized for Beckman System Gold HPLC, Pump Module 1 Type: 118, Detector Model: 166. If using other HPLC systems, some adjustment may be required. 96-well filter plate format can be adapted to automated liquid handling system. Some adjustments in procedure may be required. SDS-PAGE is a simple way to evaluate the efficiency of sample fractionation by IgY microbeads. The major plasma proteins, such as albumin, transferrin, IgG, and Apo-A1, can be easily visualized by Coomassie blue staining of SDS gel. The different protein banding patterns of neat plasma, flow-through fraction and eluted fraction represent the protein composition in each sample. Under reducing condition (with DTT or BME in SDS sample buffer), the heavy and light chains of IgG are separated and migrate at different speed; while under nonreducing conditions (no DTT or BME in SDS sample buffer), the heavy and light chains of IgG are linked by disulphide bounds and migrate on gel as single band at higher molecular weight. The total protein mass in plasma is about 60–80 mg/mL. Approximately 90% of proteins are captured by IgY-12 microbeads. The recovery rate is about 85–90%. The protein concentration in flow-through fraction is very low. To see protein banding pattern in SDS gel, load maximal volume of sample that a well of the gel can hold for flow-through and eluted/bound fractions, usually 25–30 μL(including 5× Sample Buffer). As a control, load 15–20 μL of diluted neat plasma, usually at 1 to 70–80 dilutions. After flow-through fraction is concentrated and protein concentration is measured, an equal amount of protein (5–10 μG) from each fraction and neat plasma can be loaded for comparison on SDS gel. Fractionation efficiency can be assessed. IgY microbeads can also be used to partition plasma/serum from other species, such as nonhuman primates, mouse, rat, cow, dog, etc. To ensure maximal separation efficiency, use 50% of human sample loading for other species. IgY microbeads can be recycled for at least 100 times under proper conditions. It is important to neutralize the beads immediately after stripping.

Acknowledgments The author would like to thank Dr. Wei-Wei Zhang and Mr. Robert Gans for critical reading and editing of the manuscript. The IgY-12 microbeads products were developed and manufactured by GenWay Biotech and marketed by Beckman Coulter.

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References 1. Leslie, G. A. and Clem, L. W. (1969) Phylogeny of immunoglobulin structure and function. 3. Immunoglobulins of the chicken. J. Exp. Med. 130, 1337–52. 2. Hadge, D. and Ambrosius, H. (1984) Evolution of low molecular weight immunoglobulins – IV. IgY-like immunoglobulins of birds, reptiles and amphibians, precursors of mammalian IgA. Mol. Immunol. 21, 699–707. 3. Du Pasquier, L., Schwager, J., and Flajnik, M.F. (1989) The immune system of Xenopus. Annu. Rev. Immunol. 7, 251–75. 4. Larsson, A. and Mellerstedt, H. (1992) Chicken antibodies: a tool to avoid interference by human anti-mouse antibodies in ELISA after in vivo treatment with murine monoclonal antibodies. Hybridoma 11, 33–9. 5. Larsson, A., Balow, R. M., Lindahl, T. L., and Forsberg, P. O. (1993) Chicken antibodies: Taking advantage of evolution – A review. Poultry Science 72, 1807–12. 6. Warr, G. W., Magor, K. E., and Higgins, D. A.. (1995) IgY: clues to the origins of modern antibodies. Immunol. Today 16, 392–98. 7. Schade, R. and Hlinak, A. (1996) Egg yolk antibodies, state of the art and future prospects. ALTEX. 13, 5–9. 8. Zhang, W.-W. (2003). The use of gene-specific IgY antibodies for drug target discovery. Drug Discovery Today 8, 364–71. 9. Patterson, R., Youngner, J. S., Weigle, W. O., and Dixon, F.J. (1962) Antibody production and transfer to egg yolk in chicken. J. Immunol. 89, 272–8. 10. Stuart, C. A., Pietrzyk, R. A., Furlanetto, R. W., and Green, A. (1988) Highaffinity antibody from hens’ eggs directed against the human insulin receptor and the human IGF1 receptor. Anal. Biochem. 173, 142–50. 11. Gassmann, M., Thommes, P., Weiser, T., and Hubscher, U. (1990) Efficient production of chicken egg yolk antibodies against a conserved mammalian protein. FASEB J. 4, 2528–32. 12. Larsson, A., A. Karlsson-Parra, and J. Sjoquist. (1991) Use of chicken antibodies in enzyme immunoassays to avoid interference by rheumatoid factors. Clin. Chem. 37, 411–14. 13. Fang, X., Curran, K. W., Huang, L., Xiao, W., Strauss, W., Harvie, G. Feitelson, J., and Zhang, W.-W. (2003) Polyclonal gene-specific IgY antibodies for proteomics and abundant plasma protein depletion, in frontiers of biotechnology and pharmaceuticals, Vol. 4 (Reiner, J., Zhao, K., Chen, S.-H., and Guo, M., eds.)„ Science Press USA, Inc. Monmouth Junction, NJ, pp. 222–45. 14. Fang, X., Huang, L., Feitelson, J. S., and Zhang, W.-W. (2004) Affinity separation: divide and conquer the proteome. Drug Discovery Today: Technology 1, 141–48 15. Huang, L., Harvie, G., Feitelson, J. S., Gramatikoff, K., Herold, D.A., Allen, D. L., Amunagama, R., Hagler, R. A., Pisano, M. R., Zhang, W.-W., and Fang, X. (2005) Immunoaffinity separation of plasma proteins by IgY microbeads: meeting the needs of proteomic sample preparation and analysis. Proteomics 5, 3314–28.

5 Proteomics of Cerebrospinal Fluid: Methods for Sample Processing John E. Hale, Valentina Gelfanova, Jin-Sam You, Michael D. Knierman, and Robert A. Dean

Summary Cerebrospinal fluid (CSF) provides an important source of potential biomarkers for brain disorders and therapeutic drug development. Applications of proteomic technology to the identification and quantification of proteins in CSF are increasing rapidly. Key to obtaining reproducible and reliable data about protein levels in CSF are standardization of methods for sample collection, storage, and subsequent sample processing. Methods are described here for all steps of sample processing for a number of different proteomic approaches.

Key Words: Cerebrospinal fluid; mass spectrometry; proteomics; silver staining; two-dimensional gel electrophoresis.

1. Introduction Cerebrospinal fluid is the interstitial fluid that bathes the ventricles of the brain. CSF is produced at a rate of approx 500 mL/d (1) and participates in maintenance of hydrodynamic pressure, transportation of nutrients, and removal of metabolites from the brain (2). Because of its proximity to the different regions of the brain, CSF has long been considered an important source for biomarkers of diseases of the brain. CSF is physically separated from plasma by the blood-brain barrier (bbb). This prevents the free flow of large molecules (such as proteins) from one space to the other. Smaller molecules may diffuse more freely however penetration of the bbb is dependent on the physical From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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properties of the individual molecule. The proteins of CSF have been studied by two-dimensional (2-D) gel electrophoresis for some time (3). Many of the proteins seen in CSF are abundant serum proteins, which has led to the mistaken impression that CSF is simply an ultrafiltrate of serum. There are many differences between the protein composition of CSF and serum however. Thus CSF is not a simple ultrafiltrate of serum (1). Several proteins have been identified that are present at much higher concentration in CSF than in serum (4). Proteins produced and secreted in the brain will be prevented from rapidly diffusing into the serum by the same blood brain barrier that limits diffusion of molecules into the brain. The use of CSF for biomarker discovery and measurement is of obvious importance. However, differential levels of proteins may arise for many different reasons. Of major concern is blood contamination, which may occur during sample collection. The protein concentration of blood is 200–400 times higher than that for CSF so a very small percentage of blood can have a very dramatic effect on the protein profile of CSF of some proteins may be altered by differential handling of CSF samples and care should be taken to ensure that samples are collected and stored in as similar a fashion as possible. The success of any proteomic analysis of CSF is largely dependent on the quality of the sample analyzed. Although the samples are not collected by the analytical chemist directly, an understanding of the clinical methods used for collection and the criteria for acceptance of samples is important in downstream interpretation of the results. This section is intended as a guide to aid the analytical chemist in understanding the issues involved in sample collection and in assessing the suitability of individual samples for subsequent analysis. Lumbar puncture (LP) is routinely performed to collect cerebral spinal fluid (CSF) for confirmation of suspected meningeal infection and subarachnoid hemorrhage. CSF is also frequently examined to detect malignancies, neurodegenerative processes, and other pathology involving the central nervous system (CNS). LP also provides an opportunity for direct measurement of intracranial pressure (5,6). Although gross assessment of CSF provides clues about the presence of disease, physicians rely on a broad spectrum of laboratory techniques to evaluate patient specimens. The wealth of data available from standardized chemical, cytological, and microbiological tests on CSF from healthy individuals and patients with specific diseases provides the background against which clinical laboratories and physicians compare data for individual patients. This knowledge base allows physicians to narrow down the diagnostic possibilities suggested by a patient’s complaint and presentation (5,7). In clinical research, LP has increasingly been paired with varied analytical technologies to better characterize normal and disease biology and enhance diagnostic accuracy (8,9). The procedure also is used to characterize the central

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disposition, pharmacodynamic and clinical responses produced by established and candidate therapeutics. In drug development, analyses of CSF are used to determine if drugs penetrate the CNS and alter biochemical pathways and cellular responses. This approach aims to more thoroughly define the mechanism of action and the optimal dose for drugs designed to act on the CNS (10). Collection of CSF for clinical research generally employs LP techniques similar to those used in routine clinical practice. In attempting to identify central pharmacodynamic effects, LP is occasionally performed before and following multidose administration of drug designed to achieve circulating steady state concentrations (11). Placement of an indwelling catheter for continuous sampling of CSF from the thecal sac has also been reported in clinical research (12). This approach provides an opportunity to characterize diurnal changes in CSF. Continuous CSF sampling also creates an opportunity to evaluate acute pharmacokinetic and pharmacodynamic responses with various pharmacological interventions (10). Lumbar puncture for CSF collection in research is generally safe (13). Nevertheless, LP is an invasive procedure. Whether done for diagnostic or research purposes, the procedure carries a number of inherent risks. The most common complication from LP is postdural puncture headache. The headache is typically frontotemporal and may be accompanied by neck stiffness, dizziness, and nausea. The headache may result from added tension on anchoring structures of the brain because of removal of CSF (14). Leakage of CSF at the dural puncture site may be an important factor. The latter explanation is supported by a decreased incidence of headache when LP is performed using a small gauge spinal needle shown to reduce CSF leakage in an in vitro cadaveric dural model (15). Maintaining the patient in a prone, slightly head down position can help resolve the headache and associated symptoms. Persistent headache can be treated by intravenous administration of caffeine or epidural injection of autologous whole blood (blood patch) at the LP site (14). Other more serious, but rare risks include hemorrhage, infection, and herniation of the brain. As a result, LP is contraindicated in individuals with a bleeding diathesis, thrombocytopenia (platelet count <30,000/uL), infection and congenital malformations of the lumbar puncture site, and increased intracranial pressure because of a mass lesion. Careful evaluation and preparation of the patient and close attention to the technique ensures that these more serious risks are exceptionally rare. Patient cooperation and proper positioning of the patient are important. The spinal needle passes through multiple tissues before puncture of the dura. Although, clinical circumstances may necessitate placing the patient in different positions, the positions typically employed are designed to make the subject comfortable while fully extending the space between the spinous processes of the L4–L5

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or L3–L4 interspaces through which the spinal needle will pass (5,16). If the impact of the position on the planned analyses is unknown, it may be useful to record or predetermine the position in the research protocol. Strict attention to maintenance of sterile conditions is required. Administration of a local anesthetic reduces the risk of pain and tends to reduce unwanted movement of the patient during the course of the procedure. The care taken to avoid risk to a patient also helps ensure access to a high quality CSF specimen. A traumatic LP increases the likelihood of blood in the CSF specimen. Excessive blood contamination is particularly problematic as it can distort chemical measurements and cell counts performed on CSF. If sufficiently severe, this issue can compromise clinical interpretation or research conclusions. The possibility of a traumatic tap producing a blood contaminated specimen can sometimes occur even with careful preparation of the patient and close attention to procedural technique. Accordingly, CSF is typically collected in a series of sterile tubes that are submitted for laboratory analyses in a fashion that reduces the potential for such confounds. In diagnostic settings, the first tube is typically used for chemical and immunological analyses, a second for microbiological examination, and a third for total cell count and differential cell counts (17). Despite such precautions, clinical laboratories may refuse to perform selected analyses on grossly bloody CSF specimens. Similarly, research laboratories should pay close attention to this common, undesired source of preanalytical variability and define criteria that render a CSF specimen unacceptable for the intended use. Once collected, the CSF must be processed, preserved, and transported in a timely manner so as not to render a good specimen unacceptable. Preservation of specimen constituents consistent with the state at the time of collection requires attention to the potential impact of the collection device and vials, materials used for processing time and temperature. It is important to assure that “biology in the tube” does not confound attempts to characterize the biological state under investigation through degradation of chemical and cellular elements. Although we realize we cannot anticipate all possible proteomic methodologies that may be applied to CSF, the purpose of this review is to provide basic methodology for the collection, storage, qualification, and processing of CSF for proteomic applications. 2. Materials 2.1. Sample Simpliﬁcation 1. Montage equilibration buffer, wash buffer and columns are provide with the Montage Albumin Deplete Kit (Millipore). 2. Protein G Sepharose (Amersham Biosciences).

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2.2. Gel Electrophoresis 1. Ammonium carbonate solution: 1M ammonium carbonate adjusted to pH 11 with 30 % ammonium hydroxide. 2. Reduction-alkylation cocktail: 97.5% ACN, 2% iodoethanol, and 0.5% triethylphosphine. 3. Rehydration sample buffer: 11 cm IPG strips, (pH 3–10 nonlinear) and 8–16% linear gradient Protean II gels were from Bio-Rad, Hercules, CA, USA. 4. IPG reduction buffer: 0.1M Tris-HC1, pH 6.8, 6M urea, 2M thiourea, 1% SDS, 6.4 mM dithiothreitol (DTT), 30% glycerol, and a few grains of bromophenol blue. 5. IPG alkylation buffer: 0.1M Tris-HC1, pH 6.8, 6M urea, 2M thiourea, 1% SDS, 24 mM iodoacetamide , 30% glycerol, and a few grains of bromophenol blue.

2.3. Silver Stain 1. 2. 3. 4. 5. 6.

Fixer 1: 50% MeOH, 10% Acetic acid. Fixer 2: 5% MeOH, 10% acetic acid. DTT solution: 5 mg/mL dithiothreitol. Silver solution: 0.1% AgNO3 Developer: 30 g sodium carbonate in 1 L H2 O + 500 μL 37% formaldehyde. Stopping solution: 2.3M citric acid.

2.4. In-Gel Digestion 1. Farmers reagent: 30 mM potassium ferricyanide (solution 1) and 100 mM sodium thiosulphate (solution 2). The working reagent is 1 part solution 1 and 1 part solution 2 mixed together just before use. 2. Gel reduction-alkylation reagent: 50% 0.1M ammonium carbonate solution, 48.75% acetonitrile (by volume), 1% iodoethanol, and 0.25% triethylphosphine (final pH 10). 3. Gel trypsin solution: 20 ng/μL analytical grade (Promega,Madison, WI). One vial containing 20 μg was reconstituted with 1 mL 50 mM sodium bicarbonate, pH 8. 4. Extraction solution: 100 mM NH4 HCO3 , pH 8.5. 5. Destain reagent: 50% acetonitrile, 25 mM NH4 HCO3 , pH 8.0. 6. DTT solution 2: 50 mM DTT in 100 mM NH4 HCO3 , pH 8.0. 7. Iodoacetamide solution: 100 mM iodoacetamide in 100 mM NH4 HCO3 , pH 8.0.

2.5. Processing CSF for LC/MS/MS Analysis 1. Chicken lysozyme (Sigma, St Louis, MO) is dissolved at 1 mg/mL in H2 O and stored in single use aliquots at –80°C. Working solutions are prepared by diluting at 12.5 μg/mL in urea solution before use. 2. Urea solution: 8M urea in 100 mM ammonium carbonate, pH 11.0. 3. LC/MS reduction-alkylation reagent: 97.5 % acetonitrile, 2% iodoethanol, 0.5% triethylphosphine (see Note 1).

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4. LC/MS trypsin solution: TPCK treated bovine pancreatic trypsin (Worthington, Lakewood, NJ) is dissolved at 1 mg/mL in H2 O and stored in single use aliquots at –80°C. Working solutions are prepared by diluting to 5 μg/mL in 100 mM ammonium bicarbonate pH 8.0 before use.

2.6. MALDI-TOF Mass Spectrometry 1. O-methylisourea solution: 1M solution O-methylisourea-hemisulfate (Arcos) in 100 mM sodium carbonate buffer adjusted to pH 10 with 5.0N NaOH (see Note 2). 2. Matrix solution: Recrystallized -cyano-4-hydroxycinnamic acid (see Note 3) is reconstituted with 200 μL 50 % acetonitrile, 0.05 % TFA to make saturated matrix solution.

2.7. LC-MS/MS Mass Spectrometry. 1. The C-18 reversed phase column was a Zorbax SB300 1× 50 mm (Agilent). 2. Solvent A: 0.1% formic acid (Aldrich) in water (Burdick and Jackson HPLC grade). 3. Solvent B: 50% acetonitrile, 0.1% formic acid (Aldrich) in water (Burdick and Jackson HPLC grade). 4. Solvent C: 80% acetonitrile, 0.1% formic acid (Aldrich) in water (Burdick and Jackson HPLC grade).

3. Methods 3.1. Simpliﬁcation of CSF by Removal of Abundant Proteins Similar to serum, albumin and immunoglobulin comprise more than 50 % of the protein concentration of CSF. To visualize proteins of lower abundance, it is desirable to reduce the levels of these proteins before separation or analysis. One procedure for the reduction of these proteins is described here. 1. Resuspend Protein G Sepharose beads in Montage equilibration buffer, centrifuge at 500g for 2 min, and discard supernatant. Repeat Montage equilibration buffer addition and centrifuging once. Resuspend beads pellet in equal volume of Montage equilibration buffer. 2. Dilute aliquots (50 μg protein) of CSF with Montage equilibration buffer to a volume of 300 μL. 3. To the CSF samples, add 20μL Protein G Sepharose bead suspension and agitate slowly for 1 h at RT. 4. While samples are incubating with Protein G Sepharose, rehydrate Montage columns via manufacturer specifications: add 400 μL of Montage equilibration buffer to column insert and centrifuge at 500g for 2 min. Discard eluate from the collection tube. 5. Pellet Protein G Sepharose beads at 500g for 2 min.

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6. Carefully remove (without disturbing beads) and transfer 280 μL of the effluent to a rehydrated Montage column. Centrifuge column at 500g for 2 min. Reapply the eluate to the column and centrifuge again. Add two consecutive 100-μL washes of Montage wash buffer over the column via 500g centrifugation for 2 min (final volume approx 500 μL). Discard the column insert. 7. Speed vacuum samples to approx 30–50 μL. Monitor volumes during speed vacuuming to prevent evaporation to dryness.

3.2. Processing CSF for 2-Dimensional Gel Electrophoresis Separation of proteins by 2-dimensional gel electrophoresis may be adversely affected by the presence of salts in the sample. Additionally, smearing of bands has been noted in 2D gel electrophoresis that is attributable to incomplete reduction of disulfides in proteins (18). We have addressed both of these issues in the procedure that follows. 1. Dialyze CSF against deionized water using 3-kDa MWCO dialysis tubing. 2. To reduce and alkylate the protein samples, place 50 μg of CSF protein (approx 50 μL) in a tube. Add 5 μL of ammonium carbonate solution followed by 50 μL of reduction-alkylation cocktail ( see Note 1). Cap the tube and incubate for 60 min at 37°C. 3. The sample is then uncapped and evaporated in a speedvacuum with medium heat for 2 h. After the sample is dried, reconstitute it in Bio-Rad rehydration sample buffer. 4. Samples are isoelectrically focused using 11 cm IPG strips, (pH 3–10, nonlinear). Rehydrate the IPG strips overnight at room temperature. The IPG strips are run at 8,000 V for 60,000 Vh using an IEF cell. 5. The second dimension separation is performed in 8–16% linear gradient Protean II gels at 120 V for 2 h. 6. Stain the gels with silver (Section 3.3) and scan with a Bio- Rad FX-Imager at 50-μm resolution.

3.2.1. Alternative 2-Dimensional Gel Electrophoresis Procedure This procedure incorporates the alkylation of cysteines into the prepping of the IPG strips for the second dimension. 1. After dialysis against water, dissolve samples in approx 0.4 mL of rehydration solution and apply to immobilized pH gradient strips. The rehydration of the IPG strips is performed overnight at room temperature. Run the rehydrated IPG strips at 8,000 V for 60,000 Vh using an IEF cell. 2. Following focusing, reduce the proteins on the strips by immersing the strips in IPG reduction buffer for 15 min at room temperature. 3. Alkylate the proteins by immersing the strips in IPG alkylation buffer for 15 min at room temperature.

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4. The IPG strips are then overlayed on SDS PAGE gels (8–16 % linear gradient Protean II gels) and separated at 120 V for 2 h.

3.3. Mass Spectrometry Compatible Silver Stain Some silver staining procedures use reagents, such as glutaraldehyde, which will interfere with proteomic analysis of proteins (19). Glutaraldehyde will react with lysines in proteins and cross-link them. This interferes with trypsin, which is the typical enzyme used for in-gel digestion procedures. Formaldehyde is capable of inducing the reduction of silver while not cross linking proteins. The following procedure has been used for staining of gels before in-gel digestion and mass spectral analysis. 1. Following electrophoresis soak the gels in fixer 1 for 30 min (see Note 4). Then for an additional 30 min in fixer 2. Agitate on a rocker platform gently. 2. Rinse gels with Milli-Q H2 O with gentle agitation for 20 min to 1 h (200 mL at a time). 3. Soak gels in a DTT solution for 30 min (volume sufficient to cover gel i.e., 100 mL) with gentle rocking. 4. Pour the DTT solution off and add 100 mL silver solution to cover gel which is soaked 30 min with gentle rocking. 5. Pour off the silver solution and rinse the gel once with 100 mL H2 O (rapidly). Rinse the gel twice with 100 mL developer for ∼30 sec. Then soak in 100 mL developer until proteins reach the desired intensity (see Note 5). 6. Stop the development by adding 5 mL stopping solution per 100 mL developer and agitating for 10 min. 7. Rinse the gel with several changes of distilled H2 O over 30 min. Fig. 1 shows an image of a silver stained 2-D gel of albumin depleted CSF. Fig. 2 lists the spots that were identified after digestion and LC/MS/MS analysis.

3.4. In-Gel Digestion Procedure 3.4.1. Standard Protocol Following gel separation and staining, a common procedure for identification of proteins is excising the gel band or spot and digestion of the protein in the gel with a proteolytic enzyme (most commonly trypsin). The following procedure may be used with dye or silver stained gels. 1. Place Coomassie blue or Sypro ruby-stained gel pieces in Eppendorf centrifuge tubes or in the wells of a 96-well plate. 2. Silver stained gel pieces are first de-stained by immersing them in Farmers reagent for 5–10 min followed by 3 washes with Milli-Q water (see Note 6).

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Fig. 1. Albumin was removed from a pooled CSF sample using the procedure in Section 3.1 without the protein G step. Proteins were separated by two-dimensional electrophoresis. The gel was silver stained. 3. Immerse the gel pieces in gel reduction-alkylation reagent sufficient to completely cover them (usually ∼100 μL (see Note 1). Cap the tubes, or seal the plate, and incubate the samples at 37°C for 60 min. 4. After incubation, the reagent is drawn off and discarded. Dehydrate the gel pieces with 100 μL of pure acetonitrile for approx 5 min. 5. Decant the excess acetonitrile, and dry the gel pieces on a speed vacuum for at least 15min with medium heat. 6. Rehydrate the gel pieces with gel trypsin solution (usually 5–10 μL,) and incubate overnight at 37°C (see Note 7). 7. For LC/MS/MS analysis, extract peptides with extraction solution for 60 min at 37°C and desalt with a μC-18 Ziptip before injection onto the mass spectrometer. For MALDI analysis, extract peptides as described in Section 3.6.

3.4.2. Alternative Procedure for Reduction and Alkylation of Gel Pieces For proteins separated in gels without prior reduction and alkylation this procedure can be substituted for the reduction/alkylation step described previously.

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Fig. 2. Spots were cut out and digested with trypsin (see 3.4). LC-MS/MS was performed with the procedure listed in Section 3.7 and proteins identified searching the non-redundant database with SEQUEST.

1. Crush excised Coomassie blue or Sypro stained gel pieces and soak in destain reagent for at least 5 min. Decant destain and add fresh destain reagent for another 5 min. Repeat at least 3 times total. 2. Silver stained gel pieces are destained as described previously. 3. Reduce protein in the gel with 50 μL of DTT solution 2, for 30 min at 56°C. 4. Remove the DTT solution and replace with 50 μL of iodoacetamide solution, for 30 min at 45°C. 5. Remove the iodoacetamide solution, wash the gel pieces with extraction solution, and then with acetonitrile and dry on a speedvacuum. 6. Rehydrate the gel pieces with gel trypsin solution (usually 5–10 μL,) and incubate overnight at 37°C (see Note 7). 7. For LC/MS/MS analysis, extract peptides with extraction solution for 60 min at 37°C and desalt with a μC-18 Ziptip before injection onto the mass spectrum. For MALDI analysis, extract peptides as described in Section 3.6.

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3.5. Processing CSF for LC/MS/MS Analysis More frequently, gel electrophoresis procedures are being eliminated and protein mixtures are being analyzed by chromatographic separation of proteolytic digests of the mixtures. This procedure has been used for processing of unfractionated cerebrospinal fluid before LC/MS/MS analysis. 1. Prepare the albumin and immunoglobulin depleted CSF as in Section 3.1. 2. Add 40 μL of chicken lysozyme solution (see Note 8). 3. Add 100 μL of LC/MS reduction/alkylation reagent (see Note 1). Close the tubes and incubate for 1 h at 37°C. 4. Speedvacuum the solutions to dryness, at least 3 h with medium heat. 5. Redissolve the pellet in 200 μL of LC/MS trypsin solution to produce a 1.6M urea solution and an enzyme: substrate ratio of 1:50 (w/w). 6. Incubate at 37°C overnight. 7. Typically, 100 μL of this digest is injected onto the mass spectrometer.

3.6. MALDI-TOF Mass Spectrometry Peptides from in-gel digests of proteins may be identified from their peptide fingerprint. MALDI-TOF mass spectrometry can provide that fingerprint which consists of the singly protonated mass of each tryptic peptide in a given protein. These fingerprints may be used to search protein databases using software written for this purpose. Conversion of lysines into homoarginines increases the sensitivity of detection of lysine containing tryptic peptides (20–22). This procedure has been used for 1 and 2 D gel separated proteins. 1. Following in-gel trypsin digestion (Section 3.4), extract peptides with Omethylisourea solution for 60 min at 37°C to convert lysines to homoarginine 2. Desalt samples with a μC-18 Ziptip before MALDI mass spectral analysis. 3. Spot the desalted peptides onto a MALDI target (1 μL) and add an equal volume of saturated matrix solution. 4. After the spots dry, MALDI-TOF spectra are obtained on a Voyager DE-Pro mass spectrometer (see Note 9). 5. Search peptide masses using the program Knexus. The mass of the homoarginine residue (170.23) and the S-ethanol-cysteine residue (147.03) are added to the list of user defined modifications. For peptides obtained from in-gel digests, the oxidized methionine modification is also selected.

3.7. LC-MS/MS Mass Spectrometry For unfractionated protein digests or for gel digests that contain multiple proteins, LC-MS/MS analysis can separate and fragment peptides providing

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spectra that can be used to search databases for peptide identification. The following procedure is capable of separating peptides from digests of unfractionated CSF. 1. Inject tryptic digests (100–200 L from Section 3.5) onto a C-18 reversed phase column at a flow rate of 50 L/min on a Surveyor HPLC system (ThermoFinnigan). 2. The gradient conditions are 90% solvent A, 10% solvent B to 5% solvent A, 95% solvent B over 120 min, followed by a 0.1 min ramp to 100% solvent C, and held at 100% solvent C for 5 min, followed by a 0.1 min ramp to 90 % solvent A, 10% solvent B, and held for 17 min. Divert the effluent to waste for the first 5 min to keep the source clean. The total column effluent is connected to the electrospray interface of an LTQ ion trap mass spectrometer (ThermoFinnigan). The source is operated in positive ion mode with 4.8 kV electrospray potential, a sheath gas flow of 20 arbitrary units, and a capillary temperature of 225°C. The source lenses are set by maximizing the ion current for the 2+ charge state of angiotensin. 3. Collect data in the triple play mode with the following parameters: centroid parent scan set to 1 microscan and 50 ms maximum injection time, profile zoom scan set to 3 microscans and 500 ms maximum injection time, and a centroid MS/MS scan set to 2 microscans and 2000 ms maximum injection time. Set dynamic exclusion settings to a repeat count of one, exclusion list duration of two minutes, and rejection widths of –0.75 m/z and +2.0 m/z. 4. Carry collisional activation out at a relative collision energy of 35% and an exclusion width of 3 m/z. 5. Search MS/MS spectra against a nonredundant protein database with SEQUEST (23).

4. Notes 1. The reduction alkylation solution should be prepared just before use. Triethylphosphine is pyrophoric and should be handled in a fume hood in accordance with the material safety data sheet. 2. If the pH is not properly adjusted to 10.0 the reaction may not proceed to completion. 3. -cyano-4-hydroxycinnamic acid (Sigma, St Louis, MO) is recrystalized with the following procedure. Place 50 mg -cyano-4-hydroxy-cinnamic acid in a 50 mL tube and add 10 mL of DI water. Add 30 % ammonium hydroxide until completely dissolved. Centrifuge and pour into a new 50-mL tube. In a fume hood add neat trifluoroacetic acid until pH is below 3 to precipitate the -cyano4-hydroxycinnamic acid (check with indicator paper), centrifuge, and decant the supernatant. Wash the pellet 3× with 0.1 % TFA. Dissolve the final pellet into enough 50% acetonitrile 0.1% TFA to just dissolve it, no more than 40 mL. Aliquot 200 μL into 1.0 mL tubes, speed vaccuum to dryness and store at –20°C.

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4. To minimize background staining, clean glassware with which the gel will come into contact with 50% HNO3 . Wear gloves when handling the gel as fingerprints will produce background stain. 5. Development should be executed with continuous agitation. If a precipitate should begin to form discard the developer and replace it with fresh developer. 6. The gel pieces should be clear. If yellow color remains continue washes until it is gone. 7. The volume of trypsin solution added should be just enough to rehydrate the gel. There should not be excess liquid immersing the gel. 8. Lysozyme is added as an internal standard to monitor the efficiency of reduction/alkylation and digestion. 9. The mass spectrometer was calibrated with des-Arg-bradykinin, angiotensin I, glu1-fibrinopeptide,and ACTH [1–17, 18–39, and 7–38], insulin (bovine), thioredoxin (Escherichia coli), and apomyoglobin (horse) (Sigma, St. Louis, MO).

References 1. Huang, T., Chung, H., Chen, M., Giiang, L., Chin, S., Lee, C., Chen, C., and Liu, Y. (2004) Supratentorial cerebrospinal fluid production rate in healthy adults: quantification with two-dimensional cine phase-contrast MR imaging with high temporal and spatial resolution. Radiology 233:603–8. 2. Nilsson C., Lindvall-Axelsson M., and Owman C. (1992) Neuroendocrine regulatory mechanisms in the choroid plexus-cerebrospinal fluid system. Brain Research - Brain Res. Rev. 17(2):109–38. 3. Rohlff C. (2000) Proteomics in molecular medicine: applications in central nervous systems disorders. Electrophoresis 21(6):1227–34. 4. Walsh M. J., Limos L., and Tourtellotte W. W. (1984) Two-dimensional electrophoresis of cerebrospinal fluid and ventricular fluid proteins, identification of enriched and unique proteins, and comparison with serum. J. Neurochem. 43(5):1277–85. 5. Ebers, G. Lumbar puncture and cerebrospinal fluid analysis. In: Kelley’s Textbook of Internal Medicine, 4th ed. Humes, H. D., ed. Lippincott Williams & Wilkins, Philadelphia. 2000, pp 2996–98. 6. Griggs, R. C., Józefowicz, R. F., Aminoff, M. J. Approach to the patient with neurologic disease. In: Cecil Textbook of Medicine, 22nd Ed. Philadelphia, Eds: Goldman L, Ausiello D. Saunders, pp 2196–2205. 7. Smith G. P., Kjeldsberg C. R. Cerebrospinal, synovial, and serous body fluids, In: Clinical Diagnosis and Management by Laboratory Methods, 19th Ed. Henry J. B., ed. W.B. Saunders Company, Philadelphia. 1996, pp 457–82. 8. Altemus M., Fong J., Yang R.,Damast S., Luine V., and Ferguson D. (2004) Changes in cerebrospinal fluid neurochemistry during pregnancy. Biol. Psychiatr. 56(6):386–92. 9. Baker D. G., West S. A., Nicholson W. E., et al. (1999)Serial CSF corticotropinreleasing hormone levels and adrenocortical activity in combat veterans with posttraumatic stress disorder. Am. J. Psychiatr. 156(4):585–8.

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10. Bieck P. R.and Potter W. Z. (2005) Biomarkers in psychotropic drug development: integration of data across multiple domains. Ann. Rev. Pharmacol. Toxicol. 45, 227–46. 11. Boz C., Ozmenoglu M., Velioglu S., Kilinc K., Orem A., Alioglu Z., Altunayoglu V.. (2006) Matrix metalloproteinase-9 (MMP-9) and tissue inhibitor of matrix metalloproteinase (TIMP-1) in patients with relapsing-remitting multiple sclerosis treated with interferon beta. Clin. Neurol. Neurosurg. 108:124–28, 12. Geracioti T. D., Jr, Orth D. N., Ekhator N. N., Blumenkopf B., Loosen P. T. (1992) Serial cerebrospinal fluid corticotropin-releasing hormone concentrations in healthy and depressed humans. J. Clin. Endocrinol. Metab. 74:1325–30 13. Jhee S. S., Zarotsky V. (2003) Safety and tolerability of serial cerebrospinal fluid (CSF) collections during pharmacokinetic/pharmacodynamic studies: 5 years experience. Clin. Res. Reg. Affairs 20(3):357–63. 14. Raskin N. H. Headache. In: Harrison’s Principles of Internal medicine, 16th ed. Kasper D. L., Fauci A. S., Longo D. L., Brauwald E., Hauser S. L., and Jameson J. L. eds. McGraw Hill, New York, pp 85–94 15. Angle P. J., Kronberg J, E., Thompson D. E., et al. (2003) Dural tissue trauma and cerebrospinal fluid leak after epidural needle puncture: effect of needle design, angle, and bevel orientation. Anesthesiology 99(6):1376–82. 16. Sandoval M., Shestak W., Sturmann K., and Hsu C. (2004) Optimal patient position for lumbar puncture, measured by ultrasonography. Emergency Radiology 10(4):179–81. 17. Kjeldsberg C. R. and Knight J. A. (1986) Body Fluids, Laboratory Examination of Amniotic, Cerebrospinal, Seminal, Serous, and Synovial Fluids: A Textbook Atlas, 2nd ed. American Society of Clinical Pathologists Press, Chicago, IL. 18. Herbert, B., Galvani, M., Hamdan, M., et al (2001) Reduction and alkylation of proteins in preparation of two-dimensional map analysis: why, when, and how. Electrophoresis 22, 2046–57. 19. Shevchenko, A., Wilm, M., Vorm, O., Mann, M.(1996) Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68, 850–58. 20. Hale, J. E.,. Butler, J. P, Knierman, M. D., Becker, G. W. (2000) Increased sensitivity of tryptic peptide detection by MALDI-TOF mass spectrometry is achieved by conversion of lysine to homoarginine. Anal. Biochem. 287, 110–17. 21. Brancia, F. L., Oliver, S. G., Gaskell, S. J. (2000) Improved matrix-assisted laser desorption/ionization mass spectrometric analysis of tryptic hydrolysates of proteins following guanidination of lysine-containing peptides.Rapid Commun. Mass Spectrom. 14, 2070–73. 22. Beardsley, R. L., Karty, J. A., Reilly, J. P. (2000) Enhancing the intensities of lysine-terminated tryptic peptide ions in matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 14, 2147–53. 23. Eng, J. K., McCormack, A. L., Yates, J. R. (1999) An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database, Anal. Chem. 71, 4981–88.

6 Sample Preparation of Bronchoalveolar Lavage Fluid Baptiste Leroy, Paul Falmagne, and Ruddy Wattiez

Summary Respiratory diseases are important health problem throughout the world. The bronchoalveolar lavage (BAL) fluid obtained by fiber-optic bronchoscopy is a biofluid reflecting the expression of secreted pulmonary proteins and the products of activated cells. The characterization of the BALF proteome provides an opportunity to establish diagnostic and get prognostic indicators of airway diseases. The main part of the chapter is devoted to the description of the most effective and reliable method of BALF sample preparation and processing for proteomic studies. Principal BALF proteome characteristics and the difficulties to work with this fluid are also introduced. Finally, the best conditions for high resolution and high reliability 2-dimensional electrophoresis (DE) of BALF samples are given.

Key Words: BALF sampling; bronchoalveolar lavage fluid; epithelial lining fluid; lung; mass spectrometry; proteome.

1. Introduction The respiratory tract, and in particular the alveoli, are covered by a thin film called epithelium lining fluid (ELF). This fluid plays important functions of protection against external aggressions and preservation of the gas-exchange capability of the airways. ELF contains several categories of cells (mainly of the immune system) and a wide variety of soluble components (lipids, nucleic acids, and proteins/peptides) (1). Diverse techniques exist to sample this particular fluid, but the most commonly used remains the bronchoalveolar lavage (BAL) during fiber-optic bronchoscopy. This minimally invasive and From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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reproducible procedure consists of liquid instillation through a fibrobronchoscope and recovery of the bronchoalveolar lavage fluid (BALF) by gentle aspiration (2,3). This biofluid allows the monitoring of the expression of secreted pulmonary proteins and the products of activated cells. In the last decade, the number of BALF focusing studies has grown exponentially. BALF proteome study has been initiated by some groups that have demonstrated that investigations of BALF proteome and the discovery of disease associated proteins should contribute to a better knowledge of the lung at the molecular level and to the study of the lung disorders at the clinical level (4–8). The most abundant proteins are plasma proteins that derive by diffusion across the blood barrier. A comparative analysis of serum and BALF proteomes revealed the presence of proteins specifically produced in the airways (9). These proteins are, therefore, good candidates for lung specific disease biomarkers. The proteins produced locally are very heterogeneous and can be classified according to their function: proteins involved in defense mechanisms, tissue remodelling, lipid metabolism, inflammatory processes, cell growth, and oxidant-antioxidant systems. Differential-display proteomics studies showed that nonphysiological conditions cause significant modifications of the BALF proteome; these modifications can be used to better understand pathogenesis mechanisms or to reveal diseases. Significant efforts have been devoted to large scale identification of BALF proteins or their degradation products and to the evaluation of potential markers for lung diseases, especially, for fibrosing interstitial lung diseases such as sarcoidosis and idiopathic pulmonary fibrosis (4,6,8,10,11). A majority of BALF proteomic studies use the two dimensional gel electrophoresis (2-DE) approach associated with mass spectrometry technologies. The technological progress encountered recently also allow scientists to reach the objective of quantitative analysis that is of crucial importance in differential-display proteomic. Reproducible two-dimensional BALF proteome patterns can be obtained using immobilized pH gradients (IPG) in the first dimension (Fig. 1) (12). After staining, the two dimensional patterns are compared to reveal up- or downregulated proteins. High throughput identification of proteins by mass spectrometry techniques also permits the creation of reference gels of the BALF proteome (9). These gels are now available on the World Wide Web (http://W3.umh.ac.be/biochim/proteomic.htm). The complexity of BALF proteome has been demonstrated by the great diversity of proteins present in this fluid. Although BALF proteome analysis is now largely facilitated by previous works, BALF sample preparation remains a critical step (13). Indeed, majors problems associated with BALF proteome analysis are ascribable to the low

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pI

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100 kDa α1-antitrypsin

Albumin

IgG heavy chain, γ SP-A

MW

Immunoglobulin light chain, κλ

Transthyretin

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CC16

Fig. 1. 2-D gel electrophoresis of human BALF proteins.

protein content and the high salt (because of the phosphate-buffered saline used for the lavage procedure) and lipid concentrations of this sample. Moreover, as for other human fluid, the wide dynamic range of protein concentration with the occurrence of some major proteins gives rise to difficulties in the study of lowabundance proteins. In this context, several efforts have been made to optimize BALF sample preparation and to identify less abundant proteins (13). Recently, alternative or supplement methods to the 2-DE approach have been developed such as multidimensional liquid chromatography (multi-LC): proteins from BALF are separated by chromatography (ionic and/or reverse phase) coupled “on line” with mass spectrometry analysis (multi LC-MS) (10,14). This new promising approach allows the identification and the quantification of small, minor, and hydrophobic proteins present in the BALF samples. Nevertheless, at this moment, LC–MS technology dedicated to BALF proteome study lacks quantitative data. In this chapter, we describe BALF sample preparation for 2-DE, the most currently used proteomic tool. Clearly, this sample preparation dedicated to 2-DE is also adequate to LC-MS approach. Finally, the conditions used in our laboratory to achieve 2-DE separation of BALF proteins are also detailed.

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2. Materials 1. Sterile saline solution: 0.9% (w/v) NaCl, filtered through a 0.2-μm syringe filter unit (Minisart, Sartorius, Germany). 2. Ultrapure water (double-distilled, deionized, >18) is used for all reagent preparations. 3. Protease inhibitor cocktail tablets (Complete mini, Roche, Germany). 4. Dialysis membrane: Spectra/Por membrane (MWCO : 3.5 kDa) (Spectrum). 5. Dialysis buffer: 50 mM NH4 HCO3 . 6. Speed Vac. 7. Immobilized pH gradient 3–10 NL 180mm (Pharmacia-Amersham). 8. Rehydration solution: 7 M urea, 2 M thiourea, 4% CHAPS (w/v), 2% ampholytes 3–10 (v/v), 65 mM DTE, and a trace of bromophenol blue. Prepare fresh each time. 9. Sample buffer: 7 M urea, 2 M thiourea, 4% CHAPS (w/v), 2% ampholytes 3–10 (v/v), 65 mM DTE and trace amounts of bromophenol blue. Prepare fresh each time. 10. Equilibration buffer: 50 mM Tris-HCl, pH 6.8, 6 M urea, 30% glycerol (w/v), 2% SDS (w/v). Prepare fresh each time. 11. DryStrip Reswilling tray (Amersham biosciences or Bio-Rad). 12. Multiphor II (Amersham biosciences) or Protean IEF Cell (Bio-Rad) device. 13. Standard vertical electrophoresis units for SDS-PAGE. 14. Programmable power supply able to deliver >3000 V. 15. Thermostatic circulator (Multitemp II, Amersham biosciences).

3. Methods 3.1. BALF Sampling 1. Under medical controle, place a flexible fiberoptic bronchoscope through an endotracheal tube wedged into a subsegmental bronchus of an anesthetized patient (see Note 1 and 2). 2. Instill, through the bronchoscope, 20 mL of sterile saline solution warmed to 37°C. 3. Collect the fluid by gentle aspiration and dispose in sterile siliconated bottles on ice (see Note 3). All subsequent manipulation of the samples should be realized on ice. 4. Repeat procedure 1–3 four times and combine harvested fluids except the first sample (see Note 4). 5. To elimine cells from the sample, centrifuge the combined fluids at 800g for 5min at 4°C (see Note 5, 6, 7). 6. To avoid sample degradation, add an adequat amount (1 tablet for 60 mL of BALF sample) of protease inhibitor cocktail tablets (Complete mini, Roche, Germany) (see Note 8). 7. Store BALF at –80°C until use (see Note 9).

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3.2. BALF Preparation for 2-DE Numerous BALF sample preparations have been used by different authors. Among these, precipitation (using trichloracetic acid or two-step combination of precipitant) is one of the most commonly used methodologies for protein analysis. However, limitations are observed with such a procedure because of a nonquantitative protein resolubilization after the precipitation step (13). Finally, the most efficient method of preparing a BALF sample before 2-DE analysis uses ultra filtration and dialysis-lyophilization (see Note 10). 1. Rehydrate dialysis membranes (cut-off value 3.5 kDa) by soaking them in ultrapure water heated to about 90°C for 5 min or following manufacturer’s instructions. Length of the membrane must be adapted to the sample volume. Alternatively or if small volumes need to be dialyzed, ready to use dialysis devices are available commercially (Slide-A-Lyser 3.5K, Pierce). 2. Rinse extensively the rehydrated membranes with ultrapure water. The membranes may not run dry. 3. Close the dialysis tubes on one side and improve sealing with ultrapure water. 4. Fill the membrane with BALF sample and close the second side of the dialysis tube. Check seal. 5. Immerse the membrane tube in dialysis buffer (see Note 11) and place on a magnetic stirrer at 4°C for 3 h. 6. Replace dialysis buffer twice. 7. Harvest dialyzed BALF sample into an appropriate container. 8. Reduce BALF sample volume to 20 μL in the Speed Vac (see Note 12). 10. Suspend the sample in a minimum volume of sample buffer dedicated to 2-DE analysis or LC-MS method. 11. Centrifuge at 18,000g for 15 min at 4°C to remove any unsolubilized material. 12. Sample is now ready for protein assay and 2-DE or LC-MS analysis (see Note 13).

Proteomic analysis of BALF is often hampered by the predominance of several highly abundant proteins including albumins and immunoglobulins. Depletion of these proteins is necessary before proteome analysis for detection of minor proteins. See other chapters in this book. 3.3. BALF 2-DE analysis Here we propose the optimum conditions for obtaining high resolution and reliable 2-DE of BALF samples (see Note 14). Use of different pH gradients in IEF or reticulations in SDS-PAGE should be needed in narrower study of BALF proteins. Indeed, the high number of protein spots in the pH range 4.5–6.7 are increased or decreased in different lung pathologies such as sarcoidosis or fibrosis (6,8,9). In this context, the use of narrow range IPG strips for

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IEF improves the resolution of the separation and increases the probability of detecting less abundant proteins. 1. Place the IPG strips in an adequate DryStrip Reswilling tray. 2. Cover the IPG strips first with 500 μL of rehydration buffer and second with lowviscosity paraffin oil, and let the strips rehydrate overnight at room temperature. 3. Remove the rehydrated IPG gels from the grooves, rinse them with ultrapure water, and place them, gel-side down, on water saturated filter paper. Filter papers are first blotted to remove excess water. 4. Place rehydrated IPG strip (pH 3–10, 18 cm) in the Multiphor (Amersham Biosciences) or Protean IEF Cell (Bio-Rad) device set to 20°C. The correct settings of strips and cup loading can be achieved by following the manufacturer’s instructions. 5. Apply 100 μg of protein/strip in cups at the anodic side of the IPG strip (see Note 15, 16). 6. Increase voltage linearly from 300–5,000 V during the first 3 h and stabilize the voltage at 5,000 V for 20 h (see Note 17). 7. After electrofocusing, place the IPG strips individually in capped glass tubes and equilibrate them for 20 min at room temperature in 10 mL of equilibration buffer containing 2% DTE (w/v) under gentle agitation. 8. Replace buffer by 10 mL of fresh equilibration buffer containing 2% iodoacetamide (w/v) and incubate for 20 min at room temperature under gentle agitation. 9. Run the second dimension on a 9–16% polyacrylamide linear gradient gel (18 × 20 × 1.5 cm) at 40 mA/gel constant current and 10°C. 10. The gels are then ready to be stained using standard silver staining (see Note 18).

4. Notes 1. Human bronchoalveolar lavage is an invasive method that must be carried out under the informed consent of the concerned subjects and approved by a competent ethics committee. 2. Human bronchoalveolar lavage requires topical lidocaine anaesthesia and endotracheal tube manipulation and must thus be performed by a competent physician in an adequate environment. 3. Typically, the mean recovery of BALF is 55 % of the instilled volume. 4. The first sample is separated from the others to avoid bronchial contamination. 5. With this procedure, typical protein concentration of BALF ranges from 0.05 to 1.20 mg/mL. 6. During the BAL process, the cellular elements can secrete a variety of components. Therefore, the cells should be removed immediately to provide optimal proteome stability. 7. After centrifugation, the phenotype of cells (macrophages, lymphocytes) can be analyzed. The normal cellular pattern of BALF contains mainly alveolar

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8. 9. 10.

11. 12. 13.

14.

15.

16. 17. 18.

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macrophages, a small percentage of lymphocytes and less than 2% of polymorphs. This cellular pattern changes in many lung pathologies. Protease inhibitors are necessary but peptide inhibitors, e.g., high concentration of aprotinin, may interfere with mass spectrometry analysis. Samples should be divided into small aliquots before storage to avoid repeated freezing and thawing of samples. Desalting by dialysis and ultra membrane centrifugation are very effective techniques for salt removal, leading to minimal sample loss compared to precipitation or filtration methods. However, during membrane ultracentrifugation, protein adsorption onto the membrane surface is a problem that can be, in part, circumvented by repeated washing steps with sample solution after centrifugation (13). Bath dialysis volume must be 100× sample volume. Speed Vac centrifugation is an easier method that generates less protein loss than freeze-drying (13). If electrophoresis is not to be run at this time, store the lyophilized BALF at –80°C until used. Protein concentration of BAL fluid must be determined using a suited protein assay. IEF running conditions depend on the pH gradient and the length of the IPG gel strip used. Conditions presented here assume the use of nonlinear wide-range immobilized pH gradient (3–10) 18 cm long IPG strips (optimized for body fluids) and the Multiphor II electrophoresis system of Amersham Biosciences. We also recommend the second dimension to be run on linear gradient polyacrylamide gels (9–16%) for best resolution. Different methods can be used to apply BALF sample on IPG strips such as the sample cup method or during the strip rehydration process. Classically, to increase the loading capacity, and enhance the resolution of 2-DE, the entire IPG gel can be used for sample application, with the proteins entering the gel during rehydration. Nevertheless, the best BALF 2-DE quality is obtained using the cup loading approach. A recent new type of in-gel sample application named “paper bridge sample application” has been optimized for the BALF proteome analysis. This procedure allows increasing the loading capacity of BALF sample without loss of the 2-DE resolution (8). After IEF but before equilibration, strips may be stored at –80°C until the second dimension. The quantity may be varied according to the sensitivity of the staining method. Quantification of proteins is a major problem of the 2-DE approach, especially after silver staining. However, a new fluorescent protein labeling protocol (2DDIGE) before electrophoretic separation has been developed. (See other chapters in this book).

Acknowledgments R. Wattiez is Research Associate of the Belgian FNRS. The authors thanks Catherine S’Heeren for her technical assitance.

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References 1. Wattiez, R. and Falmagne, P. (2005) Proteomics of bronchoalveolar lavage fluid. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 815, 169–78. 2. Baughman, R. P. and Drent, M. (2001) Role of bronchoalveolar lavage in interstitial lung disease. Clin. in Chest Med. 22, 331–41. 3. Reynolds, H.Y. (2000) Use of bronchoalveolar lavage in humans—past necessity and future imperative. Lung 25, 271–93. 4. Wattiez, R., Hermans, C., Bernard, A., Lesur, O., and Falmagne, P. (1999) Human bronchoalveolar lavage fluid: two-dimensional gel electrophoresis, amino acid microsequencing and identification of major proteins. Electrophoresis 20, 1634–45. 5. Noel-Georis, I., Bernard, A., Falmagne, P., and Wattiez, R. (2002) Database of bronchoalveolar lavage fluid proteins. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 771, 221–36. 6. Magi, B., Bini, L., Perari, M.G., Fossi, A., Sanchez, J.C., Hochstrasser, D., Paesano, S., Raggiaschi, R., Santucci, A., Pallini, V., and Rottoli, P. (2002) Bronchoalveolar lavage fluid protein composition in patients with sarcoidosis and idiopathic pulmonary fibrosis: a two-dimensional electrophoretic study. Electrophoresis 23, 3434–44. 7. Lindahl, M., Stahlbom, B., and Tagesson, C. (1999) Newly identified proteins in human nasal and bronchoalveolar lavage fluids: potential biomedical and clinical applications. Electrophoresis 20, 3670–6. 8. Sabounchi-Schutt, F., Astrom, J., Hellman, U., Eklund, A., and Grunewald, J. (2003) Changes in bronchoalveolar lavage fluid proteins in sarcoidosis: a proteomics approach. Eur. Respir. J. 21, 414–20. 9. Noel-Georis, I., Bernard, A., Falmagne, P. and Wattiez, R. (2001) Proteomics as the tool to search for lung disease markers in bronchoalveolar lavage. Dis. Markers 17, 271–8. 10. Kriegova, E., Melle, C., Kolek, V., Hutyrova, B., Mrazek, F., Bleul, A., du Bois, R. M., von Eggeling, F. and Petrek, M. (2006) Protein Profiles of Bronchoalveolar Lavage Fluid from Patients with Pulmonary Sarcoidosis. Am. J. Resp. Crit. Care Med. 26, 1145–54. 11. Lenz, A. G., Meyer, B., Costabel, U., and Maier, K. (1993) Bronchoalveolar lavage fluid proteins in human lung disease: analysis by two-dimensional electrophoresis. Electrophoresis 14, 242–4. 12. Lenz, A. G., Meyer, B., Weber, H., and Maier, K. (1990) Two-dimensional electrophoresis of dog bronchoalveolar lavage fluid proteins. Electrophoresis 11, 510–3. 13. Plymoth, A., Lofdahl, C. G., Ekberg-Jansson, A., Dahlback, M., Lindberg, H., Fehniger, T. E., and Marko-Varga, G. (2003) Human bronchoalveolar lavage: biofluid analysis with special emphasis on sample preparation. Proteomics 3, 962–72. 14. Wu, J., Kobayashi, M., Sousa, E., Lieu, W., Cai, J., Goldman, S. J., Dorner, A. J., Projan, S. J., Kavuru, M. S., Qiu, Y., and Thomassen, M. J. (2005) Differential

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proteomic analysis of bronchoalveolar lavage fluid in asthmatics following segmental antigen challenge. Mol. Cell. Proteomics 4, 1251–64. 15. Chromy, B., Gonzales, A., Perkins, J., Choi, M., Corzett, M., Chang, B. C., Corzett, C. H., and McCutchen-Maloney, S. L. (2004) Proteomic analysis of human serum by two-dimensional differential gel electrophoresis after depletion of high-abundant proteins. J. Proteome Res. 6, 4–8.

7 Preparation of Nasal Secretions for Proteome Analysis Begona Casado, Paolo Iadarola, and Lewis K. Pannell Summary The determination of protein patterns in nasal secretions of healthy subjects can help in the early diagnosis of diseases such as acute sinusitis. The comparison of nasal lavage fluid collected from subjects with acute sinusitis before and after pharmacological treatment gives information about the drug effects on glandular secretions. Nasal secretions were stimulated with 1× NS (0.9% Normal Saline) and 24× NS in healthy subjects and in sinusitis subjects before and after pharmacological treatment. The nasal lavage fluid (NLF) proteins are precipitated with a solution of “acid-ethanol.” Using this solution, the high molecular weight proteins precipitate and separate from the low molecular weight proteins. The proteins are digested and the peptides are separated using a capillary liquid chromatographic system. Eluted peptides are analyzed on ESI-Q-TOF mass spectrometry instrument.

Key Words: CapLC-ESI-Q-ToF; liquid-liquid extraction; Nasal secretions; pharmacological treatment; proteomics; sample preparation; sinusitis.

1. Introduction Nasal secretions (NS) are a barrier against pathogenic (e.g., bacteria and viruses) and nonpathogenic (e.g., fine particles) antigens that are present in the air and are breathed in through the nose. NS serve to humidify, heat or cool, and clean inhaled air and contain proteins of the innate immune system. These proteins are from plasma, glandular mucous and serous cells (1,2) and their release is started during allergen exposure, rhinovirus, adenovirus, influenza, bacterial rhinosinusitis, cystic fibrosis, and occupational exposure. The hyperresponsiveness is a typical characteristic of inflamed mucosa and airways (3–5) although different molecular mechanisms may be involved in allergic, infectious, and nonallergic disorders. The release of nasal secretions, From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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is generally induced by spraying a solution of normal (S) and hypertonic saline (HTS). The latter, in humans, is an airway irritant of nasal mucosa by stimulating nociceptive nerves and glandular secretion. For example, substance P is released in the mucosa after a neural depolarization with local axon responses caused by hypertonic saline stimulation (6,7). HTS nasal provocations have been employed also to understand the different neural response between acute sinusitis, acute rhinitis, and the nonallergic rhinitis present in subjects with chronic fatigue syndrome (CFS) (8,9). In the last 4 years, the interest of the scientific community on nasal mucosa and nasal secretions has increased. The collection of nasal lavage, in fact, is a simple way for obtaining samples from the upper airways and may be performed using noninvasive procedures. For example, the agent responsible for the Creutzfeldt-Jakob disease can be identified “in vivo” in nasal mucosa (10), and an anthrax vaccine based on the use of an anthrax protective antigen (PA) protein carried by liposomeprotamine-DNA (LPD) is nasally dosed in mice (11). The complete knowledge of the nasal secretion constituents has not been achieved yet. It is apparent that the identification of specific mucous protein profiles may help elucidate the different mechanisms involved in host defense. To date the determination of mucus protein profiles can be achieved using a proteomic procedure. Different proteomic approaches have been used so far to analyze nasal lavage fluid (NLF). Lindahl et al. used two dimensional electrophoresis (2-DE) with matrix-assisted laser desorption/ionization (MALDI) mass spectrometry to analyze either the proteome of NLFs from subjects exposed to methyltetrahydrophthalic anhydride (MHHPA) or dimethylbenzylamine (DMBA), and that from healthy nonsmokers and smokers (12–18). Another approach has been described by Casado et al. who applied liquid chromatography (LC) with electrospray ionization (ESI) mass spectrometry to analyze NLFs of subjects affected by acute sinusitis before and after pharmacological treatment, and for the comparison of NLFs of normal subjects before and after nasal provocation (19,20). Kristiansson et al. have used the same procedure for the analysis of HHPA-(hexahydrophthalic anhydride) adducted albumin tryptic peptides in nasal lavage fluid as biomarkers of exposure (21,22). These two complementary approaches provided new information on proteins involved in host protection and defence against microorganisms and occupational exposure. 2. Materials 2.1. Pharmacological Treatment 1. Antibiotic: amoxicillin-clavulanic acid (Augmentin, GlaxoSmithKline, Research Triangle Park, NC, USA).

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2. Steroid: fluticasone propionate nasal spray (Flonase, GlaxoSmithKline, Research Triangle Park, NC, USA). 3. 2 adrenergic agonist vasoconstrictor: oxymetazoline nasal spray (Super G, Landover, MD, USA). 4. United States Pharmacopea normal saline: saline nasal spray (0.9% NaCL) (Abbott Laboratories, IL, USA).

2.2. Nasal Provocation 1. United States Pharmacopea normal saline: saline nasal spray (0.9% NaCL) (Abbott Laboratories, IL, USA). 2. Hypertonic saline 21.6% 3. Beconase AQ pump aspirator spray device (23,24) (Glaxo-Wellcome, Triangle Park, NC, USA). 4. Nasal secretion collection: 5-ounce Dixie wax-paper cup (James River Corp., Norwalk, CT, USA) or polypropylene beakers (Fisher Scientific, Fair Lawn, NJ, USA).

2.3. Nasal Lavage Fluid Preparation for Liquid Chromatography and Mass Spectrometry Analysis 1. Protein assay on 96-well micro plates using MRX Microplate Reader Instrument (Dynex Technologies, Chantilly, VA, USA). 2. Bovine albumin as standard protein for total protein assay (Sigma, MO, USA). 3. Acid-ethanol solution: 50% 0.2N acetic acid, 50% ethanol, 0.02% sodium bisulfite (25) stored at 4°C. Ethanol and acetic acid obtained from Fisher Scientific (Fair Lawn, NJ, USA), and sodium bisulfite from Mallinckrodt Laboratory Chemicals (Phillipsburg, NJ, USA). 4. Protein digestion: sequencing grade modified trypsin (Promega, Madison, WI, USA). 5. Digestion buffer: 0.1M ammonium bicarbonate (pH 7.8) (Sigma, St. Louis, MO, USA).

2.4. Liquid Chromatography and Mass Spectrometry Analysis of Nasal Lavage Fluid 1. Desalt and concentration: 35 × 0.32 mm BioBasic C18 precolumn ( Thermo Hypersil-Keystone, Bellefonte, PA, USA). 2. Peptide separation: Reverse-phase Zorbax C18 column (100 mm × 150 μm id) (Micro-Tech Scientific, Sunnyvale, CA, USA). 3. Solvent A: HPLC grade H2 O with 0.2% formic acid (Fisher Scientific, Fair Lawn, NJ, USA). 4. Solvent B: HPLC grade acetonitrile with 0.2% formic acid (Fisher Scientific, Fair Lawn, NJ, USA).

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5. Capillary LC instrument (Waters Inc, Milford, MA, USA). 6. Electrospray-Quadrupole-Time of Flight mass spectrometer (Waters Inc, Milford, MA, USA).

3. Methods 3.1. Nasal Provocation 1. Subjects’ nasal cavities need to be pre-washed with 24 sprays (100 μL each nostril) of sterile normal saline (1 × NS, 0.9% NaCl) using a Beconase AQ pump aspirator spray device. 2. Subjects gently blow out through their noses, and the lavage fluid from both nostrils into a 5-ounce Dixie wax-paper cup. This material is discarded. 3. 10 min later step 2 is repeated using 12 sprays of 1 × NS, and the lavage fluid discarded. Now the nasal provocation is performed. 4. 100 μL of 0.9% normal saline is administered separately into each nostril. 5. After 5 min, NLF is collected using 12 sprays of 0.9% NS (see Note 1). 6. The NLF must gently blow out into a cup. NLF from left and right nostrils are mixed together (first series) (see Note 2). 7. Immediately after this collection, the same subject is sprayed with hypertonic saline (HTS) (21.6% NaCl, 24 times the tonicity of NS (24 × NS). The pH of HTS (freshly prepared solution with double distilled deionized water) is 6.07 (see Note 3). 8. After 5 min, 12 puffs of NS are sprayed into each nostril. 9. The NLF must gently blow out into a cup. NLFs from left and right nostrils are mixed together (second series). 10. After pharmacological treatment on day 6, the first and second series for day 6 are collected repeating the procedure indicated in steps 1–9. 11. Lavage fluids are gently shaken to disperse mucous globules and pipetted into Eppendorf tubes. Samples are frozen at –20°C until analysis is performed.

3.2. Nasal Lavage Fluid Preparation 3.2.1. Total Protein Assay 1. Measure the total protein concentration in each sample using modified Lowry’s method (see other chapters in this book). 2. Place standard human albumin or real samples (10 μL) in triplicate in polystyrene microtiter plates, and add assay reagents. 3. Measure the optical densities (650 nm) with a microtiter-plate reader. 4. Interpolate the protein concentrations in the samples from the regression analysis of the standard curve (protein concentration in normal NLF: 1st series 878 μg/mL and 2nd series 1,700 μg/mL; protein concentration in sinusitis NLF: day 1 1st series 1,321 μg/mL and 2nd series 1,512 μg/mL, and day 6 1st series 638 μg/mL and 2nd series 725 μg/mL).

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3.2.2. Acid-Ethanol Precipitation 1. 30 μL of nasal lavage fluid are mixed with an equal volume of 50% ethanol, 50% 0.2 N acetic acid, 0.02% sodium bisulfite. 2. Leave the protein to precipitate at –20°C overnight (see Note 4). Samples are stable in acid-ethanol solution for long time if stored at –20°C. 3. Centrifuge the mixture for 30 min at 4°C. The supernatant containing endogenous peptides, lipids and sugars is discarded.

3.2.3. Protein Digestion 1. Prepare a fresh solution of 0.1 M ammonium bicarbonate pH 7.8. 2. Dissolve the protein pellet in 10 μL of 0.1 M ammonium bicarbonate pH 7.8 vortex until the protein pellet is dissolved. 3. Dissolve 25 μg of trypsin in 25 μL of ammonium bicarbonate buffer (see Note 5). 4. Trypsin is added to the samples in trypsin:protein ratio of 1:20 (w/w). 5. Incubate the solution at 37°C overnight. 6. Inactivate the trypsin by adding 1 μL of 0.1% formic acid.

3.3. Analyis of Nasal Lavage Fluid Preparations by Liquid Chromatography Coupled to Mass Spectrometry Electrospray has the advantage of ionizing macromolecules in a liquid. The ions observed are formed by addition of proton (hydrogen ion) to give the [M+H] ion in which M = analyte molecule, H = hydrogen ion. For large macromolecules (such as peptides) there will often be a distribution of many charge states. 1. Same amounts of tryptic peptides are injected into a capillary liquid chromatography (CapLC) system after testing the LC system (see Note 6). 2. Peptide mixture are concentrated and desalted on a BioBasic C18 precolumn applying an isocratic procedure (95% water in 0.2% formic acid (FA)) with a flow rate of 20 μL/min for 10 min (see Note 7). Table 1 m/z, charge state, mean and SD of elution time, and CV of ﬁve chosen tryptic peptides from albumin from NLF. m/z 682.38 812.41 693.82 671.83 575.32

Charge state

Time (min) (X+±)

CV (%)

+3 +2 +2 +2 +2

47±0.9 61±0.9 43±1.2 47±1.3 34±0.9

1.8 1.5 2.8 2.8 2.6

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Fig. 1. Total ion chromatogram (TIC) profile of nasal lavage fluid.

Fig. 2. The histograms showing the proteins tabulated according biological function and origin, found in sinusitis NLFs pre- (Day1) and post- (Day6) pharmacological treatment.

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3. The peptides are separated on a Zorbax C18 column (100 mm x 150 μm I.D.) using a gradient from 95% water in 0.2% FA to 95% acetonitrile in 0.2% FA over 100 min. The flow is set at 10 μL/min. A splitter is used to carry 1 μL/min on the analytical column (see Note 8). 4. Separated peptides are analyzed using Electrospray-Quadrupole-Time of Flight mass spectrometer. The samples are run in duplicate. The reproducibility of elution times is determined by comparison with the retention time of five tryptic peptides of endogenous albumin (Table 1). Fig. 1 shows a total ion chromatogram (TIC) profile of nasal lavage fluid (see Note 9).

Table 2 Keratins from normal NLF are reported. We detect type I and type II reﬂecting the spectrum of cutaneous, transitional, and type I and II form heterodimers in intermediate ﬁbers respiratory mucosal cells. Sinusitis had a more limited spectrum with k1, k5, k6a, k6f, k10, and k13 Keratin k1–k2 and k9–k10 k6, k4, k16 k1–k10 k25 k6, k16 k5–k14 k8–k18, k7, k19

k5–k14 from k5–k14 to k1–k10

k6, k16 or (k17)

k5–k14

Epithelium epidermis terminally differentiated squamous cells hairs inner root sheaths outer root sheath’s inner layer outer layer and sebaceous glands transitional cuboidal epithelium and pseudoatratified respiratory epithelium basal cells (progenitors of respiratory epithelium differentiation of pseudostratified respiratory epithelial suprabasal cells wet stratified squamous epithelial lining epithelial invaginations of submucosal glands and ducts myoepithelial cells surround submucosal glands

Location anterior nares anterior nares nasal vestibule

nasopharynx

oral and esophageal mucoseae

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3.4. Data Interpretation 1. The protein identification can be performed using MASCOT MS/MS ion search software (see Note 10). 2. If the spectra are manually sequenced, the new peptide sequence can be matched to a protein using peptide match program in the protein identification resource (PIR, www.pir.georgetown.edu). 3. PIR BLAST similarity search is used to search unknown protein query sequences. 4. Protein sequence alignments are constructed using the CLUSTAW program in PIR. 5. The identified proteins are compared on the basis of their origin and function (26). In Fig. 2, the proteins identified before and after pharmacological treatment were grouped according the biological function and origin. All the inflammatory proteins were identified only on Day 1 and not on Day 6. We can hypothesize that the treatment was successful and blocked the influx of inflammatory cells (e.g., IL-16 and IL-17E), the generation of their mediators (e.g., TGF- 2 receptor), vascular permeability, and glandular hypersecretion. On Day 1, keratins associated with respiratory epithelium and the squamous metaplasia present in sinusitis were detected. Different actin protein (actin , 1, and 2) reflect the hyperplasia of the respiratory epithelium. Keratin proteome in normal NLF reflected the anticipated normal type of basal, pseudostratified, respiratory, glandular, and stratified nonkeratinized and keratinized squamous epithelium that is present in the nose. A large number of keratins have been detected in normal NLF then in sinusitis NLF. Keratin profile in sinusitis NLF was consistent with desquamation of terminally differentiated cells and the presence of squamous metaplasia. The results demonstrate the changes that are taking place in respiratory epithelial cells during inflammation (Table 2).

4. Notes 1. The subjects pressed their left nostrils closed, and then spritzed 12 sprays of 1× NS into their right nostrils. 2. Because the amount of nasal secretions blown out from left and right nostrils are different, it is important to mix the two samples together. Because both nostrils are not always simultaneously closed or open, this situation may cause a different pattern of proteins. It is very important to mix the samples as, mixing the specimens from the two nostrils, the sample homogeneity is strongly improved. Big globules of mucus must be dissolved to liberate the proteins in the mucous net. 3. Hypertonic saline stimulates the glandular secretions, local mucosal substance P release and pain. Because it is a provocation, the presence of a physician is recommended. 4. Acid-ethanol solution precipitates high molecular weight proteins. Supernatant contains peptides that can be used for following determinations. Although

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5. 6.

7.

8.

9.

10.

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proteins precipitate in few hours, it is recommended to carry out the procedure overnight. If the trypsin stock is not entirely used, we suggest resuspending it in acetic acid to prevent autodigestion of trypsin. Before injecting the NLF samples, it is important to test the LC system. one microliter of nine peptide mixtures (neurotensin, angiotensin I, angiotensin 2, Glu-fibrinopetide B, somatostatin, bradykynin, bombesin, enkephalin, and substance P; 1 pmol of each peptide) is injected on LC system. Resolution and sensitivity can be checked. During peptide concentration and desalting, small (3–4 amino acids) and highly hydrophilic peptides are washed off from the precolumn. Those losses can prevent damage of the column. It is important to look carefully at the samples before injecting them on the CapLC to check if samples are contaminated with mucous globules. This material in fact can block the capillary causing the damage to the system. For LCESI-MS/MS formic acid replaces trifluoroacetic acid (TFA) in the LC mobile phase because an efficient ionization is prevented by the strong ion pairing characteristics of TFA. Duplicate and triplicate runs are necessary to examine the reproducibility of the elution. Two options are available for checking consistency of the chromatographic system. First, it is possible to spike the sample with one or more standard peptides. Second, it is possible to choose tryptic peptides from an endogenous protein present in the sample. You must know before analyzing your sample which abundant protein is present in the sample and if it is easily cleaved by the enzyme you are using. Those peptides need to be consistently present in your runs. We choose the second option and we looked at five peptides from albumin. The mean, standard deviation of retention time, and CV % of the five peptides are calculated to see how reproducible the experiments are. The program can be found in the web (www.matrixscience.com). There is a disadvantage in using MASCOT in the web. Only the first 300 peptides can be searched on the database. Using the nonrestricted MASCOT more information can be retrieved from the raw data. The following general search parameters were used: monoisotopic molecular masses, enzyme trypsin, peptide tolerance of ± 0.4 Da and MS/MS tolerance of ± 0.3 Da. The search is restricted to Homo sapiens species to make the search easier.

References 1. Baraniuk, J. N. (2000) Immunology and Allergy Clinics of North America (Lasley, M. and V. Altman, L. C. eds.), Saunders, Philadelphia, pp. 245–64. 2. Baraniuk, J. N., Staevska, M. (2004) Current Therapy in Allergy Immunology and Rheumatology (Lichtenstein, L. M., Busse, W. W. and Geha, R. S., eds.), Mosby, Philadelphia, pp. 17–24.

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3. Meyer, P., Andersson, M., Persson, C.G., and Greiff, L. (2003) Steroid-sensitive indices of airway inflammation in children with seasonal allergic rhinitis. Pediatr. Allergy Immunol. 14, 60–65. 4. Dahl, R., and Mygind, N. (1998) Mechanisms of airflow limitation in the nose and lungs. Clin. Exp. Allergy 28, 17–25. 5. Svensson, C., Andersson, M., Greiff, L., and Persson, C.G. (1998) Nasal mucosal end organ hyperresponsiveness. Am. J. Rhinol. 12, 37–43. 6. Baraniuk, J. N., Ali, M., Yuta, A., Fang, S-Y., and Naranch, K. (1999) Hypertonic saline nasal provocation stimulates nociceptive nerves, substance P release, and glandular mucus exocytosis in normal humans. Am. J. Respir. Crit. Care Med. 160, 655–62. 7. Sanico, A. M., Philip, G., Lai, G. K., and Togias, A. (1999) Hyperosmolar saline induces reflex nasal secretions, evincing neural hyperresponsiveness in allergic rhinitis. J. Appl. Physiol. 86, 1202–10. 8. Baraniuk, J. N., Clauw, D. J., and Gaumond, E. (1998) Rhinitis symptoms in chronic fatigue syndrome. Ann. Allergy Asthma Immunol. 81, 359–65. 9. Fukuda, K., Straus, S. E., Hickei, I., Sharpe, M. C., Dobbins, J. C., and Komaroff, A. (1994) The chronic fatigue syndrome: a comprehensive approach to its definition and study. Ann. Intern. Med. 121, 953–9. 10. Tabaton, M., Monaco, S., Cordone, M. P., Colucci, M., Giaccone, G., Tagliavini, F., and Zanusso, G. (2004) Prion deposition in olfactory biopsy of sporadic Creutzfeldt-Jakob disease. Ann Neurol. 55, 294–6. 11. Sloat, B. R., and Cui, Z. (2005) Strong mucosal and systemic immunities induced by nasal immunization with anthrax protective antigen protein incorporated in liposome-protamine-dna particles. Pharm Res. Dec 6; [Epub ahead of print] 12. Lindahl, M., Stahlbom, B., and Tagesson, C. (1995) Two-dimensional gel electrophoresis of nasal and bronchoalveolar lavage fluids after occupational exposure. Electrophoresis 16, 1199–1204. 13. Lindahl, M., Stahlbom, B., Svartz, J., and Tagesson, C. (1998) Protein patterns of human nasal and bronchoalveolar lavage fluids analyzed with two-dimensional gel electrophoresis. Electrophoresis 19, 3222–29. 14. Lindahl, M., Stahlbom, B., and Tagesson, C. (1999) Newly identified proteins in human nasal and bronchoalveolar lavage fluids: potential biomedical and clinical applications. Electrophoresis 20, 3670–76. 15. Lindahl, M., Svartz, J., and Tagesson, C. (1999) Demonstration of different forms of the anti-inflammatory proteins lipocortin-1 and Clara cell protein-16 in human nasal and bronchoalveolar lavage fluids. Electrophoresis 20, 881–90. 16. Ghafouri, B., Stahlbom, B., Tagesson, C., and Lindahl, M. (2002) Newly identified proteins in human nasal lavage fluid from non-smokers and smokers using twodimensional gel electrophoresis and peptide mass fingerprinting. Proteomics 2, 112–20. 17. Lindahl, M., Stahlbom, B., and Tagesson, C. (2001) Identification of a new potential airway irritation marker, palate lung nasal epithelial clone protein, in

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20. 21.

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24. 25. 26.

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human nasal lavage fluid with two-dimensional electrophoresis and matrix-assisted laser desorption/ionization-time of flight. Electrophoresis 22, 1795–1800. Lindahl, M., Irander, K., Tagesson, C., and Stahlbom, B. (2004) Nasal lavage fluid and proteomics as means to identify the effects of the irritating epoxy chemical dimethylbenzylamine. Biomarkers 9, 56–70. Casado, B., Pannell, L. K., Viglio, S., Iadarola, P. et al. (2004) Analysis of the sinusitis nasal lavage fluid proteome using capillary liquid chromatography interfaced to electrospray ionization-quadrupole time of flight- tandem mass spectrometry. Electrophoresis 25, 1386–93. Casado, B., Pannell, L. K., Iadarola, P., Baraniuk, J.N. (2005) Identification of human nasal mucous proteins using proteomics. Proteomics 5, 2949–59. Kristiansson, M. H., Lindh, C. H., and Jonsson, B. A. (2003) Determination of hexahydrophthalic anhydride adducts to human serum albumin. Biomarkers 8, 343–59 Kristiansson, M. H., Lindh, C. H., and Jonsson, B. A. (2004) Correlations between air levels of hexahydrophthalic anhydride (HHPA) and HHPA-adducted albumin tryptic peptides in nasal lavage fluid from experimentally exposed volunteers. Rapid Commun Mass Spectrom. 18, 1592–8. Ali, M., Maniscalco, J., and Baraniuk, J. N. (1996) Spontaneous release of submucosal gland serous and mucous cell macromolecules from human nasal explants in vitro. Am. J. Physiol. 270, L595–L600. Baraniuk, J. N., Silver, P. B., Kaliner, M. A., and Barnes, P. J. (1994) Int. Arch. Allergy Immunol. 103, 202–8. Baraniuk, J. N., Okayama, M., Lundgren, J. D., Mullol, M. et al. (1990) Vasoactive intestinal peptide in human nasal mucosa. J. Clin. Invest. 86, 825–31. Wu, H. C. H., Huang, H., Yeh, Lai-Su, L., Barker, C. W. (2003) Protein family classification and functional annotation. Comput. Biol. Chem. 27, 37–47.

8 Preparation of Urine Samples for Proteomic Analysis Rembert Pieper

Summary Reproducible procedures for the preparation of protein samples isolated from human urine are essential for meaningful proteomic analyses. Key applications are the discovery of novel proteins or their modifications in the human urine as well as protein biomarker discovery for diseases and drug treatments. The methodology presented here features experimental steps aimed at limiting protein losses because of organic solvent precipitation, effective separation of proteins from other compounds in the human urine and molecular weight-based enrichment of proteins in two distinct fractions. Urinary proteins are separated from cellular debris in the urine via centrifugation, concentrated with 5-kDa-cutoff membrane concentration devices and separated via size exclusion chromatography into fractions with a higher and a lower molecular weight than 30 kDa, respectively. A successive optional affinity removal step for highly abundant plasma proteins is described. Finally, buffer exchange steps useful for specific downstream proteomic analysis experiments of urinary proteins are presented, such as 2-dimensional gel electrophoresis, differential protein or peptide labeling and digestion with trypsin for LC-MS/MS analysis.

Key Words: Biomarker discovery; gel electrophoresis; human urine; multidimensional liquid chromatography; proteomic sample preparation; urinary proteome.

1. Introduction Human urine plays a central role in clinical diagnostics. The human urinary proteome has been investigated particularly in the context of renal and bladder malfunction and cancer (1–6). Under normal physiological conditions, small protein amounts are excreted with the urinary fluid (0.5–5 mg per voiding), because the kidney restricts passage of plasma proteins, particularly in the From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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Mr range above 40 kDa during the filtration process in the glomeruli. More protein is lost in diseases, particularly those affecting the kidney, leading to proteinurea (7). Marshall and Williams as well as Anderson et al. pioneered the research in the characterization of the human urinary proteome in the eighties and nineties including sample preparation methods to isolate proteins from other matter in the urinary fluid (2,5,8–13). The most frequently used urine sample preparation methods for proteomic analysis are based on selective protein precipitation (12–17) or ultrafiltration and molecular weight-based enrichment steps (3,11,15,16,18). Urinary sample preparation should be performed at 4°C and in the presence of protease inhibitors to avoid protein degradation. Cellular debris in urinary fluid should be removed before protein enrichment to avoid contamination with cellular proteins. Urine concentration is required to effectively separate proteins in size exclusion chromatography (SEC) experiments into protein fractions of distinct Mr ranges. Immunoaffinity subtraction (IAS) permits the selective removal of highly abundant plasma proteins in urine concentrates and enrichment of lower abundance urinary proteins (3,19). Concentrated or lyophilized urinary protein samples are eventually prepared in buffers compatible with a variety of proteomic analysis techniques.

2. Materials 2.1. Urinary Protein Concentrate Preparation 1. 250-mL conical bottom polypropylene centrifugation tubes (Fisher Scientific). 2. CompleteTM protease inhibitor cocktail tablets (Roche, Indianapolis, IN). 3. Centricon® Plus-80 (5,000 NMWL) centrifugal filter devices (Millipore, Billerica, MA). 4. Amicon® Ultra-4 (5,000 NMWL) centrifugal filter devices (Millipore, Billerica, MA). 5. Swinging bucket rotor with 250-mL conical tube adaptors and centrifuge for velocities up to 4,000g (Beckman-Coulter, Fullerton, CA). 6. Buffer A: 100 mM sodium phosphate, pH 7.0, 150 mM NaCl, 0.02% sodium azide. 7. BCA assay reagents (Pierce Chemicals, Rockford, IL).

2.2. Size Exclusion Chromatography of Urinary Protein Concentrates 1. HiLoad 16/60 Superdex 75 prep grade column (GE Healthcare, Piscataway, NJ). 2. Liquid chromatography system (FPLC) with fraction collector adjustable to 4°C. 3. Centricon® Plus-20 (5,000 NMWL) centrifugal filter devices (Millipore, Billerica, MA). 4. Broad range gel filtration standard (Bio-Rad, Hercules, CA) (see Note 1). 5. Buffer B: 25 mM ammonium bicarbonate, 1 mM benzamidine, 1 mM Na-EDTA.

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2.3. Immunoafﬁnity Subtraction of Proteins 1. Multiple Affinity Removal Spin Cartridge for the Depletion of High Abundance Proteins in Human Serum (Agilent Technologies) or Vivapure Anti-HSA/IgG Removal Kit (Sartorius AG) or Affinity Depletion Cartridge for Removal of HSA and Immunoglobulins from Human Serum (Applied Biosystems). 2. Elution buffer: 0.5% glycine, 0.25% CHAPS, 150 mM NaCl, 2M urea, pH 2.5. 3. Ultrafree® -CL 0.45-μm centrifugal filter devices, (Millipore, Billerica, MA)

2.4. Final Sample Preparation for Proteomic Analysis 1. IPG rehydration solution: 8M urea, 2M thiourea, 4% CHAPS, 18 mM DTT and 0.5% Bio-Lyte® pH 3–10 carrier ampholytes (Bio-Lyte® is from Bio-Rad, Hercules, CA). 2. Freeze-dry/lyophilization unit (vacuum pump, evacuable centrifuge, cold trap).

3. Methods An overview of sample preparation and fractionation steps for urinary proteins is provided in Fig. 1. The schematic also shows downstream applications for urinary proteome analysis. 3.1. Urinary Protein Concentrate Preparation 1. The urine sample is collected, e.g., from a patient in a clinical laboratory, and transferred into a 250-mL tube with a conical bottom. It is cooled on ice and two CompleteTM protease inhibitor cocktail tablets are added to minimize protein degradation. The sample tube is centrifuged at 3,000g for 60 min at 4°C (see Note 2). In order not to disturb the precipitate, the supernatant is pipeted carefully into a new polypropylene tube. It can be frozen and stored for days at –80°C or processed immediately. The precipitate containing cellular debris and other insoluble matter is discarded. 2. The supernatant is transferred to a Centricon® Plus-80 device and spun at 3,000g at 4°C, until the urine sample volume is reduced to approx 4–5 mL (see Note 3). This sample is collected and transferred into an Amicon® Ultra-4 centrifugal filter device. It is concentrated to 1 mL by spinning at 4,000g at 4ºC, rediluted with buffer A to 4 mL and reconcentrated to approx 500 μL. 3. The protein concentrate is transferred to a 1.5-mL microtube and usually has a brownish color. It can be frozen and stored for days at -80°C or processed immediately. 4. The urinary protein concentrate is spun at 10,000g for 15 min at 4°C. The supernatant of the centrifugation step is recovered and the pellet discarded. The protein amount is measured and the sample is ready to be subjected to the size exclusion chromatography experiment.

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Fig. 1. Overview of urinary protein sample preparation procedures. 1. Removal of precipitates via centrifugation at 3,000g; 2. Concentration of urine in Centricon® Plus80 centrifugal filter devices; further concentration in Amicon® Ultra-4 centrifugal filter devices; 3. Size exclusion chromatography (Superdex 75) generating two fraction pools (SEC ≤30 kDa fraction and >30 kDa fraction); 4. Reconcentration of SEC samples; 5. Immunoaffinity subtraction generating the IAS >30 kDa fraction; 6. Final sample concentration in Amicon® Ultra-4 centrifugal filter devices. Downstream applications for proteomic analysis: 2-DE gel electrophoresis; digestion with trypsin (followed by LC-MS/MS analysis); LC separation of urinary proteins; covalent (isotope-coded) labeling of urinary proteins for differential quantitation using MS methods. 5. For protein quantitation using the BCA assay, 1- or 2-μL aliquots are transferred to a microtiter plate, incubated for 10 min with 100 μL BCA solution at 37ºC and measured in a spectrophotometer at = A562 . In parallel, a BCA assay standard curve with concentrations of 0.25 to 2 mg/mL bovine serum albumin is generated to calculate the total protein amount in the urinary sample.

3.2. Size Exclusion Chromatography of Urinary Protein Concentrate 1. The 16/60 Superdex 75 column is equilibrated in buffer A using an FPLC system at 4ºC in a cold cabinet (see Note 4). Once a stable baseline is observed monitoring UV light absorption at = A280 , the initial experiment pertains to the molecular

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weight (Mr ) column calibration. This experiment should be repeated on separate days to ascertain reproducibility. 2. 100 μL (approx 3.5 mg protein) of the Bio-Rad gel filtration standard (Mr range from 670 to 1.4 kDa) are loaded into the sample loop. The flow rate for the LC separation is 0.5 mL/min and fractions are eluted in volumes of approx 4–5 mL. As shown in the chromatogram of Fig. 2A, resolved LC peaks appear for the gel filtration standard proteins with the exception of the two high Mr proteins (670 and 150 kDa), which elute as a double peak. 3. Using an X/Y scatter diagram with the Mr units in logarithmic scale, the graphic display of Mr values and elution volumes should yield a nearly linear fit and enable the determination of the elution volume corresponding to the Mr of 30

Fig. 2. Size exclusion chromatography of urinary protein concentrates on a Superdex 75 column. Chromatogram A: Bio-Rad gel filtration standard with thyroglobulin (670 kDa) and Ig G (150 kDa) in double peak 1, ovalbumin (45 kDa) in peak 2, myoglobin (18 kDa) in peak 3 and vitamin B12 (1.4 kDa) in peak 4. B and C: two urinary protein concentrates, fractions 3–7: SEC >30 kDa sample pool; fractions 8–14: SEC ≤ 30 kDa sample pool. The UV280 traces were monitored. This Figure has been reproduced with permission3 .

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Pieper kDa (see Note 5). The fraction number corresponding to this elution volume is determined. After the protein concentration measurement of urinary samples using the BCA assay, the sample volume equivalent to an amount of 4 mg protein is determined. If the 500 μL urinary concentrate contains more than 4 mg protein, the appropriate sample volume is aliquoted and re-diluted to 500 μL with buffer A, while freezing the remaining concentrate. If there is less than 4 mg protein, all of the urinary protein concentrate is applied to the LC experiment. This sample is kept on ice before application to the SEC experiment. Using the same LC column, LC method, and fraction collector settings, urinary protein concentrates are loaded onto the 16/60 Superdex 75 SEC column and fractionated. As shown in the chromatograms of Fig. 2B and C for two different urinary protein samples, A280 elution traces may vary from sample to sample. The fractions should be placed on ice after collection and combined into two fraction pools: (1) the fraction pool with proteins corresponding to a Mr higher than 30 kDa (SEC >30 kDa) and (2) the fraction pool with proteins corresponding to a Mr equal to and lower than 30 kDa (SEC ≤30 kDa). The 30 kDa Mr fraction itself is added to the latter fraction pool. No fractions are collected in the baseline area (A280 = 0), usually for fractions collected before the LC peak for the 670/150 kDa gel filtration standards and after elution of the LC peak for the 1.4 kDa standard (peaks 1 and 4 in Fig. 2A, respectively). The two urinary protein sample pools are concentrated in Centricon® Plus-20 units to approx 1 mL. If fraction pool volumes are larger than 20 mL, concentrate in a stepwise process in the same Centricon® tubes. Protein amounts in the SEC >30 kDa and SEC ≤30 kDa urinary protein concentrates are measured using the BCA assay as described under Section 8.3.1. step 5 and are frozen at –80°C.

3.3. Immunoafﬁnity Subtraction of the SEC >30 kDa Fraction 1. An optional fractionation step is the enrichment of less abundant urinary proteins depleting highly abundant plasma proteins via immunoaffinity subtraction (IAS). A detailed protocol for plasma protein depletion is provided in a different chapter in this book. 2. Removal of highly abundant plasma proteins such as albumin and immunoglobulins (Ig) is desirable to increase the dynamic range for protein detection and quantitation in downstream proteomic analyses. Most abundant plasma proteins have Mr values higher than 30 kDa and are present in the SEC >30 kDa urinary fraction. This is the fraction to be subjected to IAS (see Note 6). 3. Different commercial products are available to deplete abundant plasma proteins from serum or urine and may be available in batch and/or as sealed cartridges (see Note 7). 4. This short protocol describes depletion with IAS resin in batch form. Approx 0.5–1 mL resin (available in batch or removed from a cartridge) is suspended in 1 mL buffer A and placed in a approx 5-mL microtube. The SEC >30 kDa

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fraction (500 μL) is added to the suspended IAS resin and incubated for 15 min at room temperature. During the incubation step, the suspension is occasionally agitated by gently pipeting. 5. The suspension is transferred to an Ultrafree-CL 0.45-μm filter device and centrifuged at 1,000g for 1–2 min. The filtrate (bottom tube) is collected and placed in an Amicon® Ultra-4 centrifugal filter device. The resin is resuspended gently in 1.5 mL buffer A, briefly equilibrated, spun again at 1,000g for 1–2 min and the filtrate added to the Amicon® Ultra-4 tube. This step is repeated and the three combined filtrates are concentrated to approx 250 μL by spinning the

Fig. 3. Protein spot profiles in 2-DE gels in a IAS >30 kDa fraction (top gel) and a SEC ≤ 30 kDa fraction (bottom gel). Samples with approx 200 μg protein were loaded in each IEF tube gel. In the first dimension, proteins were focused in the pI range between 4 and 7 applying 25,000 Volt-hours (Vh). The tube gel was stacked on an 8–15 %T second-dimension slab gel and proteins were resolved in the Mr range between 8 and 200 kDa over 1,300 Vh. Protein spots in the gels were stained with Coomassie Brilliant Blue G. Spot trains denoted in the gels are: (1) albumin*; (2) (-1-acid glycoprotein*; (3) Ig light chains*; (4) prostaglandin H2 D-isomerase; (5) (-1-microglobulin. *These proteins were of very high abundance in the SEC >30 kDa fraction and mostly removed via IAS. Spot trains for Ig light chains (3) and prostaglandin H2 D-isomerase (4) overlap in the bottom gel.

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Amicon® Ultra-4 device at 4,000g at 4°C. This concentrate is the IAS >30 kDa (flow-through) fraction. 6. To recycle the resin, it is resuspended in 2 mL elution buffer in the Ultrafree-CL device at room temperature, equilibrated for 2–3 min and spun at 1,000g for 1–2 min. The resin re-suspension and elution step is repeated. The eluates are discarded and the resin is immediately neutralized with 4 mL buffer A and spun at 1,000g for 1–2 min. The resin is resuspended in a small volume of buffer A (0.5–1 mL) and stored in suspension at 4°C. The resin can be reused for IAS using another urinary protein SEC >30 kDa fraction (see Note 8).

3.4. Final Urinary Protein Sample Preparation for Proteomic Analysis 1. The SEC ≤30 kDa fraction and the SEC >30 kDa fraction, if not processed via IAS, were stored at –80°C. After thawing, they are transferred to Amicon® Ultra-4 filter devices and concentrated to approx 250 μL at 4°C spinning at 4,000g, as described for the IAS >30 kDa fraction in Section 3.3. step 5. 2. Final protein amounts are determined using the BCA assay as described in Section 3.1. step 5. 3. A 40-fold buffer exchange in the Amicon® Ultra-4 follows rediluting and reconcentrating. For 250 μL, the total exchange buffer volume is therefore 10 mL, which is added stepwise to the sample in the Amicon® Ultra-4 tube (4 mL capacity). 4. For the preparation of 2-DE gel samples and for further LC separation steps, buffer B is used as the exchange buffer. The 2-DE gels in Fig. 3 display the spot profiles for two urinary protein fractions. To prepare urinary protein samples for trypsin digestion, 25 mM ammonium bicarbonate is used as the exchange buffer. These protein samples are lyophilized for 24 h and the protein amounts of the freeze-dried samples are noted. 5. For the preparation of samples in which proteins are to be covalently labeled, e.g., with isotope-coded amine-reactive tags for differential MS analysis, the exchange buffer is 50 mM HEPES, pH 7.8 and the volume is reduced to obtain a final protein concentration of approx 5 mg/mL.

4. Notes 1. The gel filtration standard contains thyroglobulin (670 kDa), bovine -globulin (150 kDa), chicken ovalbumin (45 kDa), equine myoglobin (18 kDa), and vitamin B12 (1.4 kDa) and, once reconstituted in 500 μL water, has a protein concentration of 36 mg/mL. 2. Under ideal circumstances, human urine specimens collected in clinical laboratories should be cooled on ice and supplemented with protease inhibitor tablets

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(CompleteTM ) immediately. Centrifugation at 3,000g to remove cellular debris should also occur in the clinical laboratory right after sample cooling. The resulting supernatant can be frozen at –80°C and shipped on ice. The 200–250 mL urinary fluid is added stepwise to the same Centricon® Plus-80 device (capacity of 80 mL). If precipitation occurs during the concentration step, the collected and concentrated sample is aliquoted into three 1.5-mL microtubes and spun for 15 min at 10,000g. It is ideal to equilibrate and run the Superdex 75 column at 4ºC. If a cold cabinet or a cooling system is not available, the LC experiments (gel filtration standard, urinary protein samples) can also be performed at room temperature. At room temperature, 1 mM benzamidine and 1 mM EDTA should be added to buffer A. It is important to equilibrate the reservoir with buffer A and the chromatography column to the same temperature for LC separation of the samples (either at 4ºC or at room temperature). A less ideal alternative is to approximate the 30 kDa elution volume as the fraction located equidistantly between the 45 kDa (ovalbumin) and the 18 kDa (myoglobin) LC peaks. Detailed protocols for the immunoaffinity subtraction (IAS) technology are described in a separate chapter in this book. In the human urine, several blood plasma proteins are frequently observed as highly abundant proteins (3). These proteins are human albumin (approx 67 kDa), IgG (approx 150 kDa) and -1-acid glycoprotein (approx 42 kDa). However, their concentrations are known to vary dramatically in human urine, sample- and donor-dependent fashion. Other proteins frequently of high abundance in human urine are prostaglandin H2 d-isomerase (approx 25 kDa) and -1-microglobulin (approx 31 kDa). Three simple-to-use IAS cartridges for the removal of plasma proteins are available (see Materials). Albumin and IgG are subtracted by all three immunoaffinity removal cartridges. For additional plasma protein subtraction specificities, further information should be obtained from the cartridge manufacturer. In a proteomic experiment comparing several urinary protein samples, only one of the IAS resin products should be used. The methods for the use of an IAS LC cartridge are different. Commercially available IAS LC resin cartridges (with a volume greater than 0.5 mL) are typically sufficient for high-abundance plasma protein removal from a SEC >30 kDa fraction with up to 3 mg total urinary protein. The described protocol works particularly well with protein A- or protein G-derivatized resins in which proteins A/G are cross-linked to plasma proteinspecific polyclonal antibodies via dimethylpimelimidate. In particular, this pertains to the elution and resin recycling steps which allow for effective elution of affinity-bound proteins, retain the cross-linkage and maintain the binding functions of the immobilized antibodies. Whether the elution buffer is compatible with the IAS resins of all depletion cartridges (based on cross-linkage chemistry between resin and antibodies) should be verified with the manufacturers. An alternative elution buffer provided by the manufacturer may be used for the recycling of the resin.

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References 1. Celis, J. E., Wolf, H., and Ostergaard, M. (2000) Bladder squamous cell carcinoma biomarkers derived from proteomics. Electrophoresis 21, 2115–21 2. Edwards, J. J., Anderson, N. G., Tollaksen, S. L., von Eschenbach, A. C., and Guevara, J., Jr. (1982) Proteins of human urine. II. Identification by twodimensional electrophoresis of a new candidate marker for prostatic cancer. Clin Chem 28, 160–63 3. Pieper, R., Gatlin, C. L., McGrath, A. M., et al. (2004) Characterization of the human urinary proteome: a method for high-resolution display of urinary proteins on two-dimensional electrophoresis gels with a yield of nearly 1400 distinct protein spots. Proteomics 4, 1159–74 4. Saito, M., Kimoto, M., Araki, T., et al. (2005) Proteome analysis of gelatin-bound urinary proteins from patients with bladder cancers. Eur Urol 48, 865–71 5. Williams, K. M., Williams, J., and Marshall, T. (1998) Analysis of Bence Jones proteinuria by high resolution two-dimensional electrophoresis. Electrophoresis 19, 1828–35 6. Decramer, S., Wittke, S., Mischak, H., et al. (2006) Predicting the clinical outcome of congenital unilateral ureteropelvic junction obstruction in newborn by urinary proteome analysis. Nat Med 12, 398–400 7. Waller, K. V., Ward, K. M., Mahan, J. D., and Wismatt, D. K. (1989) Current concepts in proteinuria. Clin Chem 35, 755–765 8. Anderson, N. G., Anderson, N. L., and Tollaksen, S. L. (1979) Proteins of human urine. I. Concentration and analysis by two-dimensional electrophoresis. Clin Chem 25, 1199–1210 9. Edwards, J. J., Tollaksen, S. L., and Anderson, N. G. (1982) Proteins of human urine. III. Identification and two-dimensional electrophoretic map positions of some major urinary proteins. Clin Chem 28, 941–48 10. Marshall, R. J., Turner, R., Yu, H., and Cooper, E. H. (1984) Cluster analysis of chromatographic profiles of urine proteins. J Chromatogr 297, 235–44 11. Marshall, T., and Williams, K. M. (1993) Centriprep ultrafiltration for fractionation of serum and urinary proteins before electrophoresis. Clin Chem 39, 1558 12. Marshall, T., and Williams, K. (1996) Two-dimensional electrophoresis of human urinary proteins following concentration by dye precipitation. Electrophoresis 17, 1265–72 13. Marshall, T., and Williams, K. M. (1997) Two-dimensional electrophoresis of human urine and cerebrospinal fluid following protein concentration by dye precipitation. Biochem Soc Trans 25, S657 14. Thongboonkerd, V., Chutipongtanate, S., and Kanlaya, R. (2006) Systematic evaluation of sample preparation methods for gel-based human urinary proteomics: quantity, quality, and variability. J Proteome Res 5, 183–91 15. Thongboonkerd, V., McLeish, K. R., Arthur, J. M., and Klein, J. B. (2002) Proteomic analysis of normal human urinary proteins isolated by acetone precipitation or ultracentrifugation. Kidney Int 62, 1461–69

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16. Tantipaiboonwong, P., Sinchaikul, S., Sriyam, S., Phutrakul, S., and Chen, S. T. (2005) Different techniques for urinary protein analysis of normal and lung cancer patients. Proteomics 5, 1140–49 17. Sun, W., Li, F., Wu, S., et al. (2005) Human urine proteome analysis by three separation approaches. Proteomics 5, 4994–5001 18. Oh, J., Pyo, J. H., Jo, E. H., (2004) Establishment of a near-standard twodimensional human urine proteomic map. Proteomics 4, 3485–97 19. Pieper, R., Su, Q., Gatlin, C. L., Huang, S. T., Anderson, N. L., and Steiner, S. (2003) Multi-component immunoaffinity subtraction chromatography: an innovative step towards a comprehensive survey of the human plasma proteome. Proteomics 3, 422–32

9 Isolation of Cytoplasmatic Proteins from Cultured Cells for Two-Dimensional Gel Electrophoresis Ying Wang, Jen-Fu Chiu, and Qing-Yu He

Summary Cytoplasma is the cell interior place between the cellular membrane and the nucleus, where various intracellular activities take place, including energy production, reactive oxygen species (ROS) detoxification, heme synthesis, nitrogen and lipid metabolism, phosphorylation in signal transduction, and cytoskeletal meshwork construction. The rich cytoplasmatic proteins carrying out these intracellular functions are interesting targets for biochemical and molecular biological studies. The relatively recent discipline of proteomics offers a chance to globally analyze the changes in cytoplasmic proteins corresponding to drug treatments or disease conditions, and thus provide target candidates for further biological validation in drug development and biomarker discovery. Isolation of cytoplasmic proteins from cells is a necessary step for high resolution protein separation by two-dimensional gel electrophoresis (2DE) and specific proteomic analysis.

Key Words: Cytoplasmatic proteins; protein isolation; proteomics; sample preparation; subcellular fractionation; two-dimensional gel electrophoresis.

1. Introduction The cytoplasm is crowded and highly ordered with transport vesicles, mitochondria, chloroplasts, and other organelles. The endocytosis and exocytosis in cytoplasm provide paths between the cell interior and the surrounding medium, allowing for the uptake of extracellular components and the secretion of proteins and other components produced within the cell. The cytoplasm is the place for energy metabolism (1), reactive oxygen species (ROS) detoxification (2), heme synthesis (3), nitrogen and lipid metabolism (4,5), mixed From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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phospholipids biosynthesis (6), cytoskeleton rearrangements (7), and various protein-protein interactions (8). Mapping the altered expression in cytoplasmic proteins by proteomic analysis under given conditions can provide a network of responses to intracellular and extracellular signals (9). Proteomics is a research technique that can identify, characterize, and quantitate proteins expressed in cells, tissues, or organisms under given conditions such as chemotherapeutic drug challenge (10,11). The altered proteins identified by proteomic analysis can be further characterized as potential drug targets; the global analysis of the protein alterations can result in valuable information for understanding the drug action mechanisms. By comparing the cytoplasmic protein profiles of HONE1 cells treated by gold(III) porphyrin 1a to untreated control, we identified a number of differentially expressed proteins by peptide-mass-finger printing (PMF) (12). The identification of the altered proteins provided valuable clues to illustrate the underlying drug action mechanisms.

2. Materials 2.1. Cell Culture and Wash Buffer 1. RPMI 1640 Medium or Dulbecco’s Modified Eagle’s Medium (DMEM) plus 10% FBS, supplemented with 2 mM/L l-glutamine, 100 units/mL penicillin, and 100 μg/mL streptomycin. 2. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA) (1 mM). 3. Cell washing buffer for 2DE: 10 mM Tris-HCl, pH 7.0, 250mM sucrose. 4. Teflon cell scrapers. 5. Tissue grinder.

2.2. Buffers and Reagents for Cytoplasmic Protein Precipitation 1. Extraction buffer: 10 mM Tris-HCl, pH 7.6, 10 mM KCl, 5 mM MgCl2 . 2. Nuclei isolation buffer: 10 mM Tris-HCl, pH 7.6, 10 mM KCl, 5 mM MgCl2 , 0.35M sucrose. 3. Radioimmunoprecipitation assay buffer (RIPA): 10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP 40 (w/v), 0.1% SDS (w/v), 0.5% sodium deoxycholate (w/v) (see Note 1), 1 mM DTT. 4. Protease inhibitors (see Note 2): 200 mM stock solution of phenylmethanesulfonyl fluoride (PMFS) in isopropanol (store at room temperature); 1 mg/mL leupeptin in water (store frozen in aliquots), 1 mg/mL aprotinin in water (store frozen in aliquots), 1 mg/mL pepstatin in methanol (store frozen in aliquots). 5. Phosphatase inhibitors (see Note 3): activated 200 mM stock solution of sodium vanadate in water and 200 mM sodium fluoride stock solution, store at room temperature.

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6. Methylene blue solution: 1.4% (w/v) methylene blue in 95% ethanol and filtered through 0.45 μm filter paper. 7. 10% trichloroacetic acid (TCA).

2.3. Buffers for Western Blot Analysis 1. Transfer buffer: 25 mM Tris base, 192 mM glycine, 0.05% SDS (w/v), 20% methanol. 2. TBS-T: 20 mM Tris-HCl, pH 7.6, 0.15M NaCl, 0.1% (w/v) Tween 20. 3. Stripping buffer: 50 mM glycine, 1 % SDS (w/v), pH 2.0. 4. ECL reagents (GE Healthcare).

2.4. Buffers and Reagents for Two-Dimensional Gel Electrophoresis (2DE) 1. 2DE clean up kit (GE Healthcare). 2. Rehydration solution: 8M urea, 2% CHAPS (stored frozen in aliquots). Add 0.002% bromophenol blue, 0.005% IPG buffer, and 2.8 mg/mL DTT freshly before use. 3. SDS equilibration buffer: 6M urea, 50 mM Tris-HCl, pH 8.8, 30% glycerol, 2% SDS. 4. 1.5M Tris-HCl (pH 8.8): dissolve 181.7 g Tris base into 750 mL ddH2 O, add 12N HCl to adjust to pH 8.8 and then add ddH2 O to final volume of 1 L, after that, filtered by 0.45 μm filter paper. 5. Thirty percent acrylamide gel stock solution: 30% acrylamide (w/v), 0.8% N’N’methylenebiasacrylamide (w/v), and filtered by 0.45 μm filter paper. 6. SDS running buffer: 25 mM Tris base, 0.192 M glycine, 0.1% SDS (w/v). 7. Agarose sealing solution: 0.5% agarose (normal or low-melting point), 0.002% bromophenol blue in SDS running buffer. 8. Bromophenol blue stock solution: 1% (w/v) bromophenol blue powder, 50 mM Tris-Base, filtered through 0.45 μm filter paper. 9. Silicon oil (e.g., DryStrip ® Cover Fluid from GE Healthcare).

2.5. Silver Staining Solutions 1. Fixation solution: 40% ethanol, 10% acetic acid. 2. Incubation solution: 30% ethanol, 4.1% sodium acetate (anhydrous) (w/v), 0.2% sodium thiosulfate (anhydrous) (w/v). 3. Silver nitrate solution: 0.1% silver nitrate (w/v), 0.02% formaldehyde (v/v). 4. Development solution: 2.5% sodium-carbonate (w/v), 0.01% formaldehyde. 5. Stop solution: 1.46% sodium-EDTA·2H2 O (w/v). 6. Preservation solution: 4.0% glycerol (w/v), 30% ethanol.

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2.6. Coomassie Brilliant Blue Staining Solutions 1. Fixation solution: 40% methanol (v/v) and 5% phosphoric acid (v/v). 2. Coommasie blue G-250 staining solution: 0.08% Coomassie brilliant blue G-250 (w/v) in 12% trichloroacetic acid, pH < 1.0. 3. Coomassie blue R-250 staining solution: 0.1% Coomassie brilliant blue R-250 (w/v), in 50% methanol (v/v), and 10 % acetic acid (v/v). 4. Destaining solution: 15% methanol (v/v), 10% acetic acid (v/v), 7% acetic acid (optional) (stored up to 1 month at room temperature).

3. Methods 3.1. Cell Harvest (see Note 4) 1. When the attached cells reach about 80% confluence, discard the media and wash two times with ice cold washing buffer (see Note 5), keep 1 mL washing buffer in the dish. 2. Harvest cells by prechilled cell scrapper, then transfer 1 mL of cell suspension to a clean 2.0-mL Eppendorf tube, spin down at 3,000g for 5 min (4°C), and wash two times with washing buffer, 1 mL each time. Cell pellet should be lysed immediately for extraction of cytoplasmatic proteins. 3. When suspended cells reach about 80% confluence, spin down at 3,000g for 5 min (4°C), and wash two times with ice cold washing buffer, 10 mL for each plate, and spin down to get cell pellet. Cell pellet should be lysed immediately for extraction of cytoplasmatic proteins.

3.2. Precipitation of Cytoplasmatic Proteins (see Note 6) 1. Re-suspend about 1×107 cells in 1 mL extraction buffer with protease inhibitors (0.2 mM PMSF, 5 μg/mL leupeptin, 2 μg/mL aprotinin, and 2 μg/mL pepstatin). Incubate the cells on ice for 10 min, and lysis by addition of Triton X-100 to the final concentration of about 0.3% (w/v). 2. Homogenize cells in an ice-cold tissue grinder. 30–50 passes with the grinder are recommended; however, efficient homogenization may depend on the cell type. To check the efficiency of homogenization, pipet 2–3 μL of the homogenized suspension onto a cover slide, stained with methylene blue solution and observe under a microscope. Nuclei appear as dark blue and cytoplasm as light purple blue under white field lens. If about 80% of the nuclei are not surrounded by cytoplasm, proceed to step 3. Otherwise, perform 10–20 additional passes using the tissue grinder. Excessive homogenization should also be avoided, as it can cause damage to the heavy membranes, e.g., mitochondrial membrane, which triggers release of mitochondrial components (see Note 7). 3. Slowly add 0.5 volume of nuclei isolation buffer to the bottom of extraction buffer, then remove nuclei by centrifugation at 500–700g for 10 min (swinging-bucket rotor) (4°C).

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4. Further centrifuge the supernatant from nuclei isolation at 10,000g for 30 min (fixed-angle rotor) (4°C). Save the supernatant as cytoplasmatic fraction, and the pellet as heavy membrane fraction containing mitochondria. 5. Add 0.25 volumes of 10% TCA to cytoplasmatic fraction to precipitate cytoplasmatic proteins; incubate on ice for 10 min. Spin down at 5,000g for 5 min (4°C). Remove supernatant, leaving protein pellet intact. Wash pellet with 200 μL prechilled acetone. Spin down at 5,000g for 5 min (4°C) (see Note 8). Wash twice with acetone again. Air dry the pellet, and add rehydration solution for 2DE analysis (see Note 9). Or add RIPA assay buffer for Western blot analysis to test the purity of the isolated subcellular fraction.

3.3. Western Blot Analysis to Detect the Purity of the Sample 1. Determine the purity of the isolated subcellular fractions by Western blot analysis against specific protein markers (Table 1). Separate the samples by onedimensional SDS PAGE. Table 1 Marker proteins of individual subcellular fractions. Subcellular localization

Marker protein

Database accession number

References

Mitochondria

Cox II Cox IV Cytochrome oxidase 1 VDAC MnSOD HSP60 mtHSP70 Cytochrome P450

gi gi gi gi gi gi gi gi

142786 142789 551699 340201 34707 77702086 292160 5733409

(13) (14, 15) (16) (14, 15, 17) (18, 19) (12) (20) (21)

Cytoplasma

Tubulin GAPDH -actin LDH Calpain Cytokeratin Vimentin Histone c-Jun c-Fos

gi gi gi gi gi gi gi gi gi gi

4507729 7669492 4501885 9257228 791040 1419564 62414289 3649600 20986521 29904

(22) (16, 23) (12) (20) (24) (25) (26) (18, 19) (27) (28, 29)

Nuclei

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2. Before transfer, soak PVDF membrane in 100% methanol for 1 min, then soak with transfer buffer. 3. After SDS PAGE, transfer the gel to the membrane electrophoretically, using transfer buffer. Assemble the filter paper, SDS gel and membrane into a blotting sandwich as shown in Fig. 1 (see Note 10). The current for transfer should be 0.8 times the area of the membrane in mA. 4. After transfer, incubate the membrane in 5% nonfat milk in TBS-T buffer (4°C overnight or room temperature for 2 h). 5. Wash the membrane three times with TBS-T buffer (10 min each, room temperature), followed by incubation with primary antibody, in 1% nonfat milk (4°C overnight or room temperature for 2 h). 6. Wash the membrane three times by TBS-T buffer (10 min each, room temperature), then incubate with secondary antibody in 1% nonfat milk (4°C overnight or room temperature, 45 min to 1 h). 7. Discard the secondary antibody and wash the membrane three times by TBS-T buffer (10 min each, room temperature). Develop the membrane using the ECL reagents. 8. Once a satisfactory exposure for the results has been obtained, the membrane is stripped with stripping buffer and then reprobed with another antibody. Wash the membrane twice with TBS-T buffer (10 min each, room temperature), then incubate the membrane in stripping buffer for 20 min (room temperature), wash twice with TBS-T again (10 min each, room temperature), then go to steps 4–7. Fig. 2 shows an example of Western blot analysis of specific protein markers of cytoplasmic and heavy membrane fraction from cisplatin treated HONE1 cells.

Fig. 1. Western Blot assembly.

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Fig. 2. Western blot analysis of cytoplasmatic fraction obtained from HONE1 cells treated with cisplatin for 6, 12, and 24 h.

3.4. Two-Dimensional Electrophoresis 1. Add 2.8 mg DTT, 5 μL carrier ampholytes or IPG buffer of the respective pH gradient, and 2 μL bromophenol blue to 1 mL rehydration stock solution (without DTT and IPG buffer) immediately before adding the protein samples. 2. Add protein samples to the above mentioned rehydration solution (volume recommendations are given in Table 2), vortex and spin down the samples. The recommended protein loading is given in Table 3. 3. IPG-strip sample loading by in-gel-rehydration. Pipet the sample solution into an IPG-strip holding device. Remove the protective cover from the IPG strip, position the IPG strip with the gel side down. To help coat the entire IPG strip, gently lift and lower the strip and slide it back and forth along the surface of the solution. Be careful not to trap bubbles under the IPG strip. Overlay the strips by Immobiline DryStrip® Cover Fluid (GE Healthcare) to ensure that rehydrated

Table 2 Rehydration solution volume per Immobiline DryStrip IPG Strip Length (cm) 7 13 18

Total volume per strip (μL) 125 250 340

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Table 3 Protein loads for silver and Coomassie blue staining. IPG Strip Length (cm)

pH-range

7

4–7 3–10, 3–10 NL 3–10, 3–10 NL 4–7 3–10, 3–10 NL

13 18

4.

5.

6.

7. 8.

9.

10.

Recommended protein load (μg) Silver stain Coomassie stain 25–50 50–75 50–100 200–400 100–400

50–100 50–150 100–200 400–1000 200–1000

Immobiline DryStrip gels do not dry out during electrophoresis. Finally, place the cover on the strip holder. IEF is performed according to a step-wise voltage increase procedure: rehydration at 30 V for 10–16 h, followed by 500 V and 1000 V for 1 h each, and 5,000–8,000 V for about 10 h with a total of 64,000 V hours (56,000 V hours is acceptable for 7-cm strips). After IEF, the strips can be subjected to second dimension immediately or can be kept at –70° C for several weeks. Prepare the 1.5-mm thick polyacrylamide gels. Seal the gel surface by 1-butanol, allow the gel to be polymerized for at least 30 min at room temperature or polymerize the gel overnight (see Notes 11 and 12). The strips after IEF are subjected to two-step equilibration in equilibration buffers with 1% DTT (w/v) for the first step, and 2.5% (w/v) iodoacetamide for the second step. Wash strips by SDS running buffer for three times before load onto the acrylamide gel (see Note 13). Electrophoresis conditions: set buffer circulation temperature to 10ºC and start the run at 15 mA per gel. After 30 min increase current to 30 mA per gel. After SDS page, visualize proteins with silver staining or Coomassie brilliant blue staining. For silver staining, fix the gels in fixation solution overnight, and then change to incubation solution for 30 min. After washing three times in water for 10 min each, stain the gels in silver nitrate solution for 40 min. Perform development for 15 min in development solution. Stop staining by stop solution and then wash the stained gels three times in water for 5 min each. Alternatively, visualize the gels by Coomassie brilliant blue stain. Fix the gels in fixation solution for Coomassie brilliant blue stain overnight, followed by Coommasie blue G-250 or R-250 stain for more than 12 h. Destain by destaining solution until background is acceptable. Acquire images by a suitable Image Scanner and preserve gels in preservation solution for further analysis. Fig. 3 shows 2DE pattern of cytoplasmatic proteins compared with the pattern of a whole cell protein extract.

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Fig. 3. 2DE pattern of cytoplasmatic proteins compared with the pattern of whole cell protein. One hundred μg of cytoplasmatic protein or whole cell protein from human nasopharyngeal carcinoma HONE1 cells was separated on 13-cm IPG-strips, pH 3–10, followed by SDS-PAGE (12% SDS gel), and visualized by silver staining.

4. Notes 1. Sodium deoxycholate is an ionic detergent to extract proteins. Prepare 10 % stock solution in water, protect the solution from light. Do not add sodiumdeoxycholate when preparing lysates for kinase assays. Ionic detergents can denature enzymes, causing them to lose activity. 2. Commercially available protease inhibitor cocktails can be used instead, for example, protease inhibitor cocktail (Catalog number P8340) from SigmaAldrich, protease inhibitor cocktail set I (Catalog number 539131) from Calbiochem. 3. Do not add phosphatase inhibitors when preparing lysates for phosphatase assays. 4. Wash cells intensively to avoid contamination of serum proteins in the culture medium. 5. Do not wash the cells with PBS in the last washing step, because PBS contains 150 mM sodium chloride, which will interfere with isoelectric focusing. Use instead 250 mM sucrose, 10 mM Tris-HCl, pH 7.5. 6. Always perform the isolation of cytoplasmic proteins on ice. Chill all buffers before use. Perform the isolation steps on ice to ensure the purity of subcellular fractions, because activities of most cellular proteases are inhibited at low temperature. 7. Under ideal homogenization conditions, particulate organelles such as nuclei, mitochondria, lysosomes, and peroxisomes remain intact. Golgi complexes, plasma membranes, and reticular organelles will fragment and can, at least to some extent, vesiculate. It is not possible to outline a general protocol suitable for the production of a reasonable homogenate for all kinds of cultured cells,

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Wang et al. because different conditions are required for different cell types and experimental purposes. Long centrifuge at high speed will cause difficulties in dissolving protein pellet in rehydration solution or RIPA assay buffer. 2DE clean-up kit can be used to improve the quality of 2DE results by removing interfering contaminants. Bubbles between the SDS gel and the membrane will lead to inefficient transfer. It is critical that all the glass plates for 2DE have been cleaned and rinsed extensively with distilled water. Clean the glass plates with distilled water followed by 95% ethanol to remove the acid and air-dry prior use. Prepare 12.5% second dimensional SDS gels for standard separations; prepare 8–10% gels to better separate large molecular weight proteins and 15% gels for low molecular weight proteins. For the equilibration step, add DTT and iodoacetamide to equilibration buffer just prior use.

Acknowledgments This investigation was partially supported by grants from Hong Kong University funding (No. 200511159099 to Q.Y.H.) and the Area of Excellence Scheme of the Hong Kong University Grants Committee.

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7. Favoreel H. W., Van Minnebruggen G., Adriaensen D., Nauwynck H. J. (2005) Cytoskeletal rearrangements and cell extensions induced by the US3 kinase of an alphaherpesvirus are associated with enhanced spread, Proc.Natl.Acad.Sci.U.S.A 102, 8990–5. 8. Yang T., Chaudhuri S., Yang L., Chen Y., Poovaiah B. W. (2004) Calcium/calmodulin up-regulates a cytoplasmic receptor-like kinase in plants, J.Biol.Chem. 279, 42552–559. 9. Jiang X. S., Dai J., Sheng Q. H., et al. (2005) A comparative proteomic strategy for subcellular proteome research: ICAT approach coupled with bioinformatics prediction to ascertain rat liver mitochondrial proteins and indication of mitochondrial localization for catalase, Mol.Cell Proteomics 4, 12–34. 10. He Q. Y. and Chiu J. F. (2003) Proteomics in biomarker discovery and drug development, J.Cell Biochem. 89, 868–86. 11. Wang Y., Chiu J. F., and He Q. Y. (2005) Proteomics in computer-aided drug design, Current Computer-Aided Drug Design 1, 43–52. 12. Wang Y., He Q. Y., Sun R. W., Che C. M., Chiu J. F. (2005) Gold(III) Porphyrin 1a induced apoptosis by mitochondrial death pathways related to reactive oxygen species, Cancer Res. 65, 11553–64. 13. Bidere N., Lorenzo H. K., Carmona S., Laforge M., Harper F., Dumont C., Senik A. (2003) Cathepsin D triggers Bax activation, resulting in selective apoptosisinducing factor (AIF) relocation in T lymphocytes entering the early commitment phase to apoptosis, J.Biol.Chem. 278, 31401–11. 14. Arnoult D., Gaume B., Karbowski M., Sharpe J. C., Cecconi F., Youle R. J. (2003) Mitochondrial release of AIF and EndoG requires caspase activation downstream of Bax/Bak-mediated permeabilization, EMBO J. 22, 4385–99. 15. Arnoult D., Parone P., Martinou J. C., Antonsson B., Estaquier J., Ameisen J. C. (2002) Mitochondrial release of apoptosis-inducing factor occurs downstream of cytochrome c release in response to several proapoptotic stimuli, J.Cell Biol. 159, 923–9. 16. Sanchez-Alcazar J. A., Bradbury D. A., Pang L., Knox A. J. (2003) Cyclooxygenase (COX) inhibitors induce apoptosis in non-small cell lung cancer through cyclooxygenase independent pathways, Lung Cancer 40, 33–44. 17. Vahsen N., Cande C., Briere J. J., et al. (2004) AIF deficiency compromises oxidative phosphorylation, EMBO J. 23, 4679–89. 18. Wang H., Yu S. W., Koh D. W., et al. (2004) Apoptosis-inducing factor substitutes for caspase executioners in NMDA-triggered excitotoxic neuronal death, J.Neurosci. 24, 10963–73. 19. Yu S. W., Wang H., Poitras M. F., et al. (2002) Mediation of poly(ADP-ribose) polymerase-1-dependent cell death by apoptosis-inducing factor, Science 297, 259–63. 20. Otera H., Ohsakaya S., Nagaura Z., Ishihara N., Mihara K. (2005) Export of mitochondrial AIF in response to proapoptotic stimuli depends on processing at the intermembrane space, EMBO J. 24, 1375–86.

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10 Sample Preparation of Culture Medium from Madin-Darby Canine Kidney Cells Daniel Ambort, Daniel Lottaz, and Erwin Sterchi

Summary A reproducible, standardized and simple sample preparation methodology is the key to successful two-dimensional gel electrophoresis (2-DE). This chapter describes step-bystep the sample preparation of culture medium from Madin-Darby canine kidney (MDCK) cells. Tips and tricks are given to circumvent possible pitfalls.

Key Words: Bicinchoninic acid (BCA) assay; culture medium (CM); isoelectric focusing (IEF); Madin-Darby canine kidney (MDCK); rehydration loading; two-dimensional gel electrophoresis (2-DE) ultracentrifugation; ultrafiltration .

1. Introduction Two-dimensional gel electrophoresis (2-DE), introduced by O’Farrell and Klose in 1975 (1,2), enabled separation of complex protein mixtures into individual protein species according to their net charge (pI) in the first dimension by isoelectric focusing (IEF) and in the second dimension according to their molecular mass (M r ) by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (3). In the conventional approach, IEF was performed in carrier ampholyte-generated pH gradients, which moved towards the cathode on prolonged focusing time. This “cathodic drift” phenomenon was thereafter remedied by nonequilibrium pH gradient gel electrophoresis (NEPHGE) (4) and finally eliminated with the invention of fixed immobilized pH gradients (IPG) (5–8). The development of microanalytical techniques, namely Edman sequencing (9–11) and mass spectrometry (12–14) enabled From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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identification of proteins at amounts available from a single 2-D gel. 2-DE advanced to the core technology of proteome analysis (7,8,15) and was brought from art to craft in an industrial standard. Appropriate sample treatment is the key to good results. The ideal sample solubilization procedure should result in the disruption of all noncovalently bound protein complexes and aggregates into a solution of individual polypeptides (15). Denaturation and reduction of proteins is achieved in the standard lysis buffer (O’Farrell 1975) (1) which is composed of 8–9 M urea, 2–4% CHAPS, 1% DTT or DTE and 0.8–2% carrier ampholytes. Hydrophobic proteins are better dissolved in 2 M thiourea and 7 M urea instead of 9 M urea (16). Optimized procedures for different sample types do exist (17). However, a general “Prepares them all” procedure is not available (18). Another highpriority issue is the removal and inactivation of all interfering substances: nucleic acids, lipids, salts, small ionic compounds, polysaccharides, proteases, and insoluble particles. Sample preparation for 2-DE is a very cumbersome and time-consuming task that is subject to trial and error. Because of the complex biological architecture of eukaryotic cells into organelles and large cellular structures, fractionation techniques are applied before comprehensively studying the subproteomes. In such reductionist approaches classic biochemical techniques, namely centrifugation and affinitymediated isolation using antibodies against molecular tags, are applied to enrich for subcellular fractions as reviewed by Yates 3rd (19). Beside analysis of cytosolic, organelle-specific and transmembrane proteins several investigations were aimed at identifying those proteins secreted by various cell types into the extracellular milieu or medium (20–24). This chapter describes the sample preparation of culture medium from Madin-Darby canine kidney (MDCK) cells for 2-DE (Fig. 1). The methodology is subsectioned into four parts with basic introductory information on each topic: cell culture (see Subheading 3.1.), ultracentrifugation and ultrafiltration (see Subheading 3.2.), protein quantitation by BCA assay (see Subheading 3.3.) and finally, rehydration loading and isoelectric focusing (see Subheading 3.4.).

2. Materials 2.1. Cell Culture 2.1.1. Equipment 1. BD FalconTM bulk packaged serological pipets (10 mL) (BD Biosciences, Franklin Lakes, NY, USA) 2. BD FalconTM individually wrapped serological pipets (25 mL) (BD Biosciences, Franklin Lakes, NY, USA)

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Fig. 1. Two-dimensional gel electrophoresis of Madin-Darby canine kidney cell culture supernatant (80 μg protein). First dimension: isoelectric focusing in an immobilized pH gradient (IPG) pH 3–10 nonlinear in a 24-cm-long gel strip. Second dimension: SDS-PAGE in a 12.5% gel. Silver stained. 3. BD FalconTM standard cell culture dish, standard tissue-culture treated (100 × 20 mm) (BD Biosciences, Franklin Lakes, NY, USA) 4. CELLSTAR® PP-test tubes (50 mL, sterile) (Greiner Bio-One Inc., Longwood, FL, USA) 5. Erlenmeyer flasks (250 mL) 6. Laminar air flow cabinet (Brouwer AG, Luzern, Switzerland) 7. Neubauer improved counting chamber (Assistent, Sondheim, Germany) 8. NUAIRETM US autoflow CO2 water-jacketed incubator NU-4750 (Vitaris AG, Baar, Switzerland) 9. Water bath

2.1.2. Solutions and Reagents 1. Dulbecco’s Phosphate Buffered Saline (D-PBS) (1X) (500 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA) 2. Foetal bovine serum (FBS) (E. U. approved South American origin, virus and mycoplasma tested) (500 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA) 3. Minimum essential medium (MEM) (1X) (with Earle´s salts, without l-glutamine) (500 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA) 4. Penicillin-streptomycin-glutamine (100X) (100 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA) 5. Trypsin-EDTA (1X) (100 mL) (GIBCO Invitrogen corporation, Grand Island, NY, USA)

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6. Culture medium: 1X MEM (see Note 1), 5% (v/v) FBS, 100 U/mL penicillin, 100 μg/mL streptomycin and 292 μg/mL l-glutamine. To 500 mL of MEM (one bottle) aseptically add 25 mL of FBS (see Note 2) and 5 mL of 100X penicillinstreptomycin-glutamine stock solution. Store at 4°C. Before use warm up to 37°C in a water bath. 7. Serum-free medium: 1X MEM (see Note 1), 100 units/mL penicillin, 100 μg/mL streptomycin, and 292 μg/mL l-glutamine. To 500 mL of MEM (one bottle) aseptically add 5 mL of 100X penicillin-streptomycin-glutamine stock solution. Store at 4°C. Before use warm up to 37°C in a water bath.

2.2. Ultracentrifugation and Ultraﬁltration 2.2.1. Equipment 1. Centricon® Plus-70 centrifugal filter devices (Millipore corporation, Billerica, MA, USA) 2. Centrifuge filter system (50 mL, 0.2 μm) (Costarcorporation, Cambridge, MA, USA) 3. Eppendorf centrifuge 5415R (Eppendorf AG, Hamburg, Germany) 4. Eppendorf tubes (1.5 mL, 2 mL) 5. KONTRON CENTRIKON TFT 70.38 fixed-angle rotor (KONTRON Instruments AG, Zürich, Switzerland) 6. KONTRON CENTRIKON T-2060 ultracentrifuge (KONTRON Instruments AG, Zürich, Switzerland) 7. KONTRON CENTRIKON ultracentrifuge tubes (32.5 mL) (KONTRON Instruments AG, Zürich, Switzerland) 8. Mettler AC 100 analytical balance (Mettler Instrumente AG, Greifensee, Zürich, Switzerland) 9. Sorvall RT6000D centrifuge (Kendro Laboratory Products AG, Zürich, Switzerland) 10. Sorvall H1000B swinging bucket rotor (Kendro Laboratory Products AG, Zürich, Switzerland) 11. Water bath

2.2.2. Solutions and Reagents 1. Ethylenedinitrilo tetraacetic acid disodium salt dihydrate (Na2 -EDTA 2H2 O, Titriplex® III) (GR for analysis) 2. Phenylmethylsulfonyl fluoride (PMSF) (Sigma, St. Louis, MO, USA) 3. 2-Propanol (GR for analysis) 4. Sodium hydroxide (NaOH) (pellets GR for analysis) 5. Tris(hydroxymethyl)aminomethane (Tris) (GR for analysis buffer substance)

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6. 0.5 M EDTA pH 8.0 stock solution: To make 100 mL of stock solution, dissolve 2 g of NaOH pellets in 80 mL of ddH2 O. Add 18.6 g of Na2 -EDTA-2H2 O under constant stirring at RT (see Note 3). Titrate solution to pH 8 with 5 N NaOH (liquid). Adjust to a final volume of 100 mL with ddH2 O and check pH again. Filter solution with a 0.2 μm bottle top filter. This solution can be stored at RT. 7. 0.1 M PMSF stock solution: To prepare 25 mL, dissolve 0.44 g of PMSF in 25 mL of 2-propanol (isopropanol). Warm up to 37°C in a water bath (see Note 4). Portion solution into 2 mL aliquots in 2 mL Eppendorf tubes and store at –20°C. 8. 0.2 M Tris pH 10.5 stock solution: To make 0.5 L, dissolve 12.12 g of Tris in 0.5 L of ddH2 O (see Note 5). Filter solution with a 0.2 μm bottle top filter. This solution can be stored at RT. 9. Sample solubilization buffer I: 20 mM Tris-HCl pH 9.0, 1 mM EDTA, 1 mM PMSF. To prepare 150 mL, dilute 15 mL of 0.2 M Tris pH 10.5 stock solution to a final volume of 150 mL in ddH2 O. Add 0.3 mL of 0.5 M EDTA pH 8.0 stock solution and 1.5 mL of 0.1 M PMSF stock solution under constant stirring at RT (see Note 6). Filter solution with a 0.2 μm bottle top filter. This solution should be prepared freshly just before use!

2.3. Protein Quantitation by BCA Assay 2.3.1. Equipment 1. Beaker (25 mL) 2. CELLSTAR® micro-plate (TC, sterile) (Greiner Bio-One Inc., Longwood, FL, USA) 3. Incubator 4. Vortex mixer 5. Vmax® microplate reader (Molecular Devices Corporation, Sunnyvale, CA, USA)

2.3.2. Solutions and Reagents 1. 2. 3. 4.

Albumin Standard Ampules (2 mg/mL, 10 x 1 mL) (Pierce, Rockford, IL, USA) BCATM Protein Assay Kit (Pierce, Rockford, IL, USA) BCATM Protein Assay Reagent A (500 mL) (Pierce, Rockford, IL, USA) BCATM Protein Assay Reagent B (25 mL) (Pierce, Rockford, IL, USA)

2.4. Rehydration Loading and Isoelectric Focusing 2.4.1. Equipment 1. Beaker (50 mL) 2. Centrifuge filter system (50 mL, 0.2 μm) (Costarcorporation, Cambridge, MA, USA) 3. Eppendorf centrifuge 5415R (Eppendorf AG, Hamburg, Germany) 4. EPS 3501 XL power supply (Amersham Biosciences, Uppsala, Sweden)

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5. IEF electrode strips (Amersham Biosciences, Uppsala, Sweden) 6. ImmobilineTM DryStrip Kit (Amersham Biosciences, Uppsala, Sweden) 7. ImmobilineTM DryStrip Reswelling Tray (7–24 cm) (Amersham Biosciences, Uppsala, Sweden) 8. Vortex mixer 9. MultiphorTM II horizontal electrophoresis apparatus (Amersham Biosciences, Uppsala, Sweden) 10. MultitempTM III thermostatic circulator (Amersham Biosciences, Uppsala, Sweden) 11. Multi-purpose rotator (Scientific Industries Inc., Queens Village, NY, USA) 12. Parafilm (50 × 15 m) (American National Can Company, Chicago, IL, USA) 13. Petri dishes 14. Tweezers

2.4.2. Solutions and Reagents 1. Bromophenol blue (BPB) (Bio-Rad Laboratories, Richmond, CA, USA) 2. 3-[(3-cholamidopropyl)-dimethylammonio]-1-propane sulfonate (CHAPS) (ultrapure) (USB corporation, Cleveland, OH, USA) 3. 1,4-dithioerythritol (DTE) (for biochemistry) (Merck, Darmstadt, Germany) 4. ImmobilineTM DryStrip pH 3–10 NL (IPG) (240 × 3 × 0.5 mm) (Amersham Biosciences, Uppsala, Sweden) 5. Mixed bed ion exchanger resin 6. Paraffin (Merck, Darmstadt, Germany) 7. PharmalyteTM 3–10 (for IEF) (Amersham Biosciences, Uppsala, Sweden) 8. Thiourea (puriss. p. a. ACS; ≥99% [RT]) (Fluka, Buchs, Switzerland) 9. ZOOM® urea (Invitrogen life technologies, Carlsbad, CA, USA) 10. DTE aliquots: 65 mM in 1.5 mL of sample solubilization buffer II (working solution). Portion 0.015 g (15 mg) of DTE in 1.5 mL Eppendorf tube and store at 4°C until use (see Note 7). 11. Sample solubilization buffer II (stock solution): 7 M urea, 2 M thiourea, 4% (w/v) CHAPS. To prepare 25 mL, dissolve 10.5 g of urea, 3.8 g of thiourea in 10 mL of ddH2 O under constant stirring at RT. Fill up to a final volume of 30 mL with ddH2 O (see Note 8). Add 5 g of mixed bed ion exchanger resin and stir for 10 min. Remove beads by filtration through a 0.2 μm bottle top filter. Dissolve 1 g of CHAPS, add a trace of BPB and portion solution into 1.5 mL aliquots in 1.5 mL Eppendorf tubes. Store at –20°C until use. 12. Sample solubilization buffer II (working solution): 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 1% (w/v) (65 mM) DTE, 2% (v/v) (0.8% (w/v)) Pharmalyte 3–10. To make up 1.5 mL, add 1.5 mL of sample solubilization buffer II (stock solution) to one DTE aliquot in 1.5-mL Eppendorf tube. Add 30 μL of Pharmalyte 3–10 and incubate for 15 minutes at RT with occasional vortexing until DTE is completely dissolved. This solution is prepared just before use (see Note 9).

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3. Methods 3.1. Cell Culture Mammalian renal tubular epithelium consists of at least seven different segments, complicating biochemical investigation of this heterogeneous tissue. Cultured monolayers of dog kidney (Madin-Darby canine kidney (MDCK)) cells display many typical features of renal tubular epithelia, such as brush border membrane, tight junctions and adherent junctions. MDCK strain II (26) was used to prepare samples of cell culture supernatants for 2-DE. High-quality protein samples for 2-DE are only obtained from serum-free media. Serumderived proteins heavily contaminate cell-derived secreted proteins in culture media (see Note 10). Hence it is essential to wash the cells thoroughly (twice in PBS or serum-free medium) when changing from complete culture media to serum-free conditions. 1. Harvest confluent MDCK cells by trypsinization. To five confluent 100-mm cell culture dishes add 1.5 mL of prewarmed (37°C) Trypsin-EDTA solution per dish and incubate at 37°C in a humidified incubator in an atmosphere of 5% CO2 until cells detach (see Note 11). 2. Resuspend trypsinized cells in culture medium. To each dish add 6.5–7 mL of prewarmed (37°C) culture medium and pool resuspended cells from the five dishes into one 50-mL Greiner tube. Fill up to a total volume of 40 mL with culture medium (see Note 12). 3. Seed 1.15 × 106 cells onto 100-mm cell culture dishes. Adjust final volume with prewarmed (37°C) culture medium to 9 mL per dish. Prepare a total of 18 dishes per condition (see Note 13). Incubate the cultures for about three days at 37°C in a humidified incubator in an atmosphere of 5% CO2 until cells are confluent (see Note 14). 4. Aspirate medium and wash cells twice in 4 mL of prewarmed (37°C) serum-free medium per dish (see Note 15). Add 4 mL of serum-free medium to each dish and incubate for 22 h at 37°C in a humidified incubator in an atmosphere of 5% CO2 . 5. Harvest cell culture supernatants. Pool media from 18 dishes into one 250-mL Erlenmeyer flask to give a final volume of 70–72 mL (see Note 13). Immediately proceed to ultracentrifugation and ultrafiltration (see Subheading 3.2.).

3.2. Ultracentrifugation and Ultraﬁltration Beside body fluids, such as human plasma, urine, and cerebrospinal fluid, cell culture supernatants are among the most difficult samples to be prepared for 2-D PAGE. Culture media contain the complete set of interfering substances that are incompatible with the first dimensional isoelectric focusing: insoluble particles (dead cells, cell debris), nucleic acids (from dead cells), lipids (membranes and exosomes (22)), salts (from medium see Note 1), small ionic

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compounds (amino acids from medium), phenolic compounds (Phenol red from medium) and proteases (secreted during cultivation). Therefore these contaminants are removed in a two-step purification strategy: 1) insoluble particles, nucleic acids and lipids by high-speed centrifugation; 2) salts, small ionic and phenolic compounds by ultrafiltration. Proteases are inhibited by addition of inhibitors and basic pH conditions. Alternative methods such as TCA/acetone precipitation (27) and dialysis must not be applied. Unspecific salt precipitation and loss of proteins are associated with these methods (see Note 16). 1. Add 140 μL of 0.5 M EDTA pH 8.0 and 700 μL of 0.1 M PMSF to 70 mL of culture medium in 250 mL Erlenmeyer flask and put on ice (see Note 17). 2. Transfer 4 × 17.5 mL of medium to four prechilled KONTRON CENTRIKON ultracentrifuge tubes. Adjust precise volumes with ddH2 O on an analytical balance and put tubes on ice. 3. Place precooled (4°C) KONTRON CENTRIKON TFT 70.38 fixed-angle rotor into KONTRON CENTRIKON T-2060 ultracentrifuge. Place tubes into the rotor in appropriate positions and close the lid. Ultracentrifuge for 60 min at 31,200 rpm (100,000g) at 4°C. 4. Carefully remove tubes from ultracentrifuge and put on ice. 5. Rinse Centricon® Plus-70 components consisting of cap, concentrate/retentate cup, sample filter cup and filtrate collection cup with ddH2 O to remove dust particles (see Note 18). 6. Place sample filter cup into filtrate collection cup and leave on ice. 7. Pool the medium by inverting tubes into the same sample filter cup and close. Then place assembled Centricon® Plus-70 centrifugal filter device into precooled (4°C) Sorvall H1000B swinging bucket rotor fixed in a Sorvall RT6000D centrifuge. Centrifuge for 60 min at 3,200 rpm (2,190g) at 4°C. 8. Discard the flow-through in the filtrate collection cup. Add 70 mL of prechilled sample solubilization buffer I (20 mM Tris pH 9.0, 1 mM EDTA, 1 mM PMSF) into sample filter cup (see Note 19). Centrifuge for 60 min at 3,200 rpm (2,190g) at 4°C. 9. Repeat step 8 twice. 10. After the last washing step place concentrate/retentate cup upside down onto the filtrate collection cup. Invert assembly and place back into centrifuge. Recover sample concentrate for 5 min at 2,200 rpm (1,000g) at 4°C. 11. Determine volume with a pipet. Typical final concentrate volumes are between 250 μL and 350 μL. 12. Transfer protein concentrate to a 1.5 mL Eppendorf tube and spin for 5 min at 13,200 rpm (16,100g) at 4°C to remove precipitates (see Note 20). Put samples on ice and proceed to protein quantitation (see Subheading 3.3.) or store at –20°C until use.

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3.3. Protein Quantitation by BCA Assay Quantitative determination of protein solubilized in modified lysis buffer (16) (7 M urea, 2 M thiourea, 4% CHAPS, 65 mM DTE and 2% Pharmalyte 3–10) is not possible. The Bradford assay (29) cannot be used for two reasons: 1) Coomassie Brilliant Blue G-250 binds to detergents (CHAPS) and carrier ampholytes (Pharmalyte 3–10); 2) Coomassie Brilliant Blue G-250 may not bind to protein at all under basic pH conditions (urea). The second problem may be remedied by acidification of sample with 0.1 N HCl before quantitation (30). Standard Lowry (31), Biuret (32), and BCA (bicinchoninic acid) (33) assays based on the reduction of Cu2+ to Cu+ for development of color interfere with thiol reducing agents (DTE) and thiourea. Thiourea forms complexes with copper ions. TCA/acetone precipitation (27) must not be applied in combination with any of these techniques. Protein may be lost on precipitation leading to underestimation of solubilized protein. The best solution to all problems mentioned above is quantitation of protein before solubilization in lysis buffer. In this section the BCA assay from Pierce is used to accurately quantitate protein concentration in sample solubilization buffer I (20 mM Tris pH 9.0, 1 mM EDTA, 1 mM PMSF). Although the BCA assay interferes with chelating agents (EDTA) concentrations below 10 mM are tolerated (34). 1. Prepare a set of albumin (BSA) standards in 1.5-mL Eppendorf tubes (see Table 1 for details). Use ddH2 O as diluent. Gently vortex tubes. There will be sufficient volume for two replications of each diluted standard. 2. Prepare protein samples in 1.5-mL Eppendorf tubes. Dilute 5 μL of each protein concentrate to a final volume of 100 μL in ddH2 O (1:20 dilution). Gently vortex tubes. While performing BCA assay put undiluted protein concentrates on ice (see Subheading 3.2. step 12).

Table 1 Preparation of diluted albumin (BSA) standards Vial A B C D E F

Volume of diluenta 150 25 50 75 88 98

a b

μL μL μL μL μL μL

Volume and source of BSA

Final BSA concentration

150 μL of stockb 75 μL of A 50 μL of A 25 μL of A 12.5 μL of A 2.5 μL of A

Use ddH2 O to prepare albumin (BSA) standards. The concentration of albumin standard stock solution is 2 mg/mL.

1000 750 500 250 125 25

μg/mL μg/mL μg/mL μg/mL μg/mL μg/mL

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3. Pipet 25 μL of each standard or unknown sample replicate into a microplate well. Standards are applied in duplicates, protein samples in triplicates. Use ddH2 O for blanks. 4. Prepare BCA working reagent by mixing 50 parts of BCATM Reagent A with 1 part of BCATM Reagent B in a 25 mL beaker. Mix thoroughly (see Note 21). For each standard, unknown sample or blank 200 μL of BCA working reagent is required. Include two extra replicates in your calculation. 5. Add 200 μL of BCA working reagent to each well. Gently shake microplate by hand for a few seconds. 6. Incubate microplate for 30 min at 37°C in an incubator. 7. Cool plate to RT and measure absorbance at 550 nm on a microplate reader (see Note 22). 8. Subtract the average 550 nm absorbance measurement of the blank replicates from the 550 nm absorbance measurements of all other individual standards and unknown sample replicates. 9. Prepare a standard curve by plotting the average blank-corrected 550 nm absorbance measurement (A550 , y-axis) for each albumin standard versus its amount (in μg, x-axis) in increasing order (see Note 23). 10. Use the standard curve to determine the protein amount of each unknown sample (Fig. 2). The protein concentration in unknown sample is calculated as follows: (protein amount of unknown in μg)/(volume of diluted sample in μL) × (dilution factor)= protein concentration in mg/mL (see Note 24). Typical protein concentrations are between 5 mg/mL and 7 mg/mL. 11. Portion undiluted protein concentrates into appropriate aliquots (80 μg for analytical load) in 1.5 mL Eppendorf tubes and store at –20°C until use (see Subheading 3.4.1).

3.4. Rehydration Loading and Isoelectric Focusing Traditionally, protein samples prepared in standard lysis buffer (O’Farrell 1975) (1) were loaded onto the basic end (cathode) of an isoelectric focusing (IEF) tube gel. Before sample loading the gel rods were prerun to establish a carrier ampholyte-derived pH gradient. On development of fixed pH gradients samples were applied with rubber frames or sample cups to rehydrated immobilized pH gradient (IPG) strips at the acidic or basic end (5–8) or simultaneously at both ends (35). The problem with cup-loading sample application techniques is that proteins may precipitate during the sample entry phase, which leads to horizontal streaking at the sample application point. This problem is remedied by in-gel sample rehydration where protein solubilized in lysis buffer is directly diluted with the rehydration solution used for IPG strip reswelling (36,37). Unfortunately, some proteins that are soluble in lysis buffer may be lost on dilution into rehydration solution because of lower concentrations of chaotropic agents and detergents. For simplicity protein sample preparation

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Fig. 2. Typical color response curve for albumin (BSA) standards using the BCA assay. Each point represents the mean of two replications.

and rehydration can be done all-in-one in modified lysis buffer (16) (7 M urea, 2 M thiourea, 4% CHAPS, 65 mM DTE and 2% Pharmalyte 3–10). This strategy works very well in combination with the Multiphor II horizontal flatbed isoelectric focusing system with final maximum voltage limited to 3,500 V for steady-state IEF (38). Higher voltage settings may become problematic, because zwitterionic detergent (CHAPS), reducing agent (DTE) and carrier ampholytes (Pharmalyte 3–10) heavily contribute to the current in the strip. 1. Thaw protein samples on ice (see Subheading 3.3.11). Dilute each aliquot of protein solution (80 μg for analytical load) to a final volume of 450 μL in sample solubilization buffer II (see Note 25). 2. Gently vortex tubes and solubilize protein for 60 min on a rotary shaker at RT. 3. Centrifuge for 30 min at 13,200 rpm (16,100g) at 22°C in a tabletop centrifuge to remove insoluble particles. 4. Slide the protective lid completely off the ImmobilineTM DryStrip Reswelling Tray and level the tray by turning the leveling feet until the bubble in the spirit level is centered. 5. Remove IPG strips (240 mm long, 3 mm wide ready-made ImmobilineTM DryStrips pH 3–10 NL cast on GelBond PAGfilm) from the freezer and warm up to RT.

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6. Evenly apply the entire sample-containing solution into the groove of the reswelling tray (see Note 26). 7. Peel off the protective cover sheet from the ready-made IPG strip starting at the acidic (+) end and grip the strip with tweezers at the overlapping basic plastic end (see Note 27). 8. Slowly lower the IPG strip (gel side down) onto the solution with the acidic (+) end oriented towards the number labels of the reswelling tray (see Note 26). 9. Cover the strip with 3 mL of paraffin oil (see Note 28). Repeat steps 6–9 for each sample. 10. Slide the lid onto the reswelling tray and rehydrate the IPG strips overnight at RT. 11. To remove rehydrated IPG strips sequentially from the reswelling tray, open the lid, slide the tip of tweezers along the sloped end of the slot and into the slight depression under the IPG strip. Grab the acidic (+) end of the strip with tweezers and lift the strip out of the tray. While still holding the strip, rinse it briefly with ddH2 O. Place it on a piece of damp filter paper at one edge to drain off excess liquid. Repeat procedure for each strip. 12. Set the temperature on the MultiTempTM III thermostatic circulator to 20°C. 13. Place the ceramic cooling plate in Multiphor II unit and make sure the surface is level. 14. Starting at the top of the plate near the cooling tubes pipet 1 mL of paraffin oil in a straight line onto the middle of the plate. Use the grid of the cooling plate as a guide. Then pipet 1 mL of paraffin oil on each side of the paraffin oil line (a total of 3 mL) onto the bottom of the plate. The additional 1 mL of paraffin oil on each side is evenly spread on the cooling plate to form a triangle which begins with the base line at the bottom and extends to the middle of the paraffin oil line (see Note 29). 15. Slowly lower the ImmobilineTM DryStrip tray onto the bottom of the paraffin oil triangle with the red (anodic, positively charged) electrode connection of the tray positioned at the top of the plate near the cooling tubes. 16. Connect the red and black electrode leads on the tray to the Multiphor II unit. 17. Pour 10 mL of paraffin oil into the tray at the bottom of the cooling plate. 18. Slowly lower the ImmobilineTM DryStrip aligner, 12 grooves side up, onto the bottom of the paraffin oil layer next to the black electrode. 19. Transfer the rehydrated IPG strips with tweezers to adjacent grooves of the aligner in the tray. Place the strips gel side up with the acidic (+) end at the top of the tray near the red electrode. 20. Cut one IEF electrode strip into two pieces each to a length of 110 mm. Moisten the two IEF electrode strips with deionized water (see Note 30). 21. Place the damp electrode strips across the acidic and basic ends of the aligned IPG strips.

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22. Align each electrode over an electrode strip, ensuring the marked side corresponds to the side of the tray giving electric contact (see Note 31). When the electrodes are properly aligned, press them down to contact the electrode strips. 23. Pour 100 mL of paraffin oil into the tray to cover the IPG and electrode strips. 24. Close the lid of the Multiphor II unit. Connect the leads on the lid to the EPS 3501 XL power supply and start IEF according to programmed parameters (see Note 32). 25. After IEF remove electrodes and IEF electrode strips. 26. Grip each IPG strip with tweezers at the overlapping basic plastic end, carefully remove it from the tray and rinse it with ddH2 O. 27. Place each IPG strip into a petri dish with the plastic side of the strip facing the inner wall of the petri dish and cover. Seal it with a piece of parafilm and store at –20° C until use. 28. IPG strip equilibration, SDS-PAGE and postseparation visualization techniques applied following sample preparation and IEF are not topic of this chapter. Useful tips and tricks concerning these methods can be found in the Amersham 2-D electrophoresis handbook (40). As an example the final 2-D map of MadinDarby canine kidney (MDCK) cell culture supernatant is shown in Fig. 1.

4. Notes 1. Composition of Minimum Essential Medium (MEM) (25): 264 mg/mL CaCl2 2H2 O, 400 mg/mL KCl, 200 mg/mL MgSO4 7H2 O, 6800 mg/mL NaCl, 2200 mg/mL NaHCO3 , 158 mg/mL NaH2 PO4 2H2 O, 1000 mg/mL d-glucose, 10 mg/mL Phenol red, 126 mg/mL l-arginine HCl, 24 mg/mL l-cystine, 42 mg/mL l-histidine HCl H2 O, 52 mg/mL l-isoleucine, 52 mg/mL l-leucine, 73 mg/mL l-lysine HCl, 15 mg/mL l-methionine, 32 mg/mL l-phenylalanine, 48 mg/mL l-threonine, 10 mg/mL l-tryptophan, 36 mg/mL l-tyrosine, 46 mg/mL l-valine, 1 mg/mL d-Ca pantothenate, 1 mg/mL choline chloride, 1 mg/mL folic acid, 2 mg/mL i-inositol, 1 mg/mL niacinamide, 1 mg/mL pyridoxine HCl, 0.1 mg/mL riboflavin, and 1 mg/mL thiamine HCl. 2. Before use fetal bovine serum (FBS) is heat-inactivated! Incubate one bottle (500 mL) of FBS for 30 min at 56°C in a water bath. Portion solution into 25 mL aliquots and store at –20°C. 3. The EDTA may not completely dissolve in 0.5 M stock solution below pH 8.0. Therefore titration of EDTA stock solution with a few drops of 5 N NaOH (liquid) is necessary. The solution is ready when the pale white color turns into a crystal clear solution. 4. PMSF is very toxic! Protect your eyes and skin. PMSF is not very stable in water and has a half-life of about 30 min. Hence the solution is prepared in isopropanol. PMSF is difficult to dissolve at RT; therefore the solution is warmed up to 37°C. 5. The 0.2 M Tris stock solution has a pH of 10.5–10.6. Do not titrate with HCl! The chloride ions extremely contribute to the current (heat production) during

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7. 8.

9. 10.

11.

12.

13.

14. 15. 16.

Ambort et al. isoelectric focusing (IEF). Any heat produced during IEF will cause protein precipitation and produce horizontal streaks in the final 2-D gel. The sample solubilization buffer I has a pH of 9.0–9.1. Do not titrate with HCl! The Tris serves as a positively charged ion that helps in solubilization of proteins and not to maintain constant pH (see Note 19). This solution should be prepared shortly before use because of the poor stability of PMSF in water (see Note 4). The 0.1 M PMSF stock solution should be warmed up to 37°C in a water bath. Otherwise the PMSF may precipitate! DTE is not very stable in solution. Hence it is stored as solid in small aliquots at 4°C. The final volume of 30 mL compensates for the dead volume of the magnetic stir bar in a 50-mL beaker and equals to a total volume of 25 mL. Add urea and thiourea in small portions with the help of a spatula under constant stirring at RT. Urea and thiourea will cool down the solution and decrease solubility! Therefore in between additions wait for several minutes until each small portion is fully dissolved. Do not heat urea-containing solutions above 37°C to avoid carbamylation of proteins! The sample solubilization buffer II (working solution) should be prepared freshly. Never reuse remaining buffer, better discard it! It is essential to wash the cells thoroughly (twice in PBS or serum-free medium) to remove serum proteins. Culture media heavily contaminated with fetal bovine serum (FBS) resemble human plasma! Before trypsinization confluent cells may be thoroughly washed (twice in 2–3 mL of PBS or serum-free medium) (see Note 10). Prewarm PBS and Trypsin-EDTA solution to 37°C in a water bath! To each dish treated with 1.5 mL of Trypsin-EDTA solution 6.5–7 mL of culture medium is added to give a total volume of 40 mL. Prewarm culture medium to 37°C in a water bath! In total 18 dishes per condition are prepared. The final volume of culture medium used per dish is 9 mL. Once MDCK cells reach confluence serum-free medium is added. The final volume of serum-free medium used per dish is 4 mL. This gives a total volume of 70–72 mL per condition and corresponds to the appropriate processing volume for the Centricon® Plus-70 centrifugal filter devices (see Note 18). The doubling time of MDCK strain II cells is one day. Confluence is reached after 3–4 days. Alternatively, use PBS instead of serum-free medium if costs are a major concern. TCA/acetone (27) may precipitate calcium-phosphate and small amino acids from the culture media. Very high contaminant concentrations may be achieved that extremely interfere with isoelectric focusing. Dialysis must not be used! Very high solute volumes are needed to remove salts and unspecific protein binding to the dialysis membrane may occur.

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17. The 0.1 M PMSF stock solution is warmed up to 37°C in a water bath. PMSF is added before putting medium on ice to avoid precipitation. EDTA inhibits metalloproteases by chelation of free metal ions. PMSF inhibits serine proteases and some cysteine proteases. Inhibitor cocktails (for example Complete Mini, EDTA-free from Roche) must not be used! These cocktails contain protein- and peptide-based inhibitors that can reach very high concentrations on ultrafiltration (up to 300X) and hence abundantly mask the secreted proteins present in the medium. 18. Usage guidelines for Centricon® Plus-70 centrifugal filter devices are given in the user guide (28). The filter material is made of a polyethersulfone Biomax membrane with a 5 kDa cut-off. 19. The pH of 9.0 from Tris serves two functions: 1) it maximizes protein extraction at basic pH conditions (almost any protein is in deprotonated state); 2) minimizes protease activity. The Tris itself serves as a positively charged ion that helps in solubilization of proteins (see Note 6). Very basic proteins may be lost! 20. On concentration protein precipitation may occur! 21. When Reagent B is first added to Reagent A, turbidity is observed that quickly disappears on mixing to yield a clear, green color. 22. Alternatively, wavelengths from 540–590 nm may be used with this method (34). 23. Amount of albumin standards (see Table 1) used: F, 0.625 μg; E, 3.125 μg; D, 6.25 μg; C, 12.5 μg; B, 18.75 μg, and A, 25 μg. The standard amount is referred to a volume of 25 μL. Average blank-corrected 550 nm absorbance values above 0.8 must not be used for standard curve preparation! 24. For example: (protein amount of unknown is 7 μg)/(volume of diluted sample is 25 μL) × (dilution factor is 20) = 5.6 mg/mL. 25. A final volume of 450 μL is recommended by the supplier (Amersham Biosciences) for rehydration of one 24 cm ImmobilineTM DryStrip. For first trial prepare a duplicate! The upper limit is twelve samples per run. 26. Avoid trapping of air bubbles. 27. Do not wear gloves during removal of protective cover sheet! The rubber material tends to stick to the “naked” gel and hence will damage it. 28. Overlaying of IPG strips with paraffin oil reduces risk of urea crystallization during rehydration. 29. In this case the paraffin oil evenly distributes the heat produced during IEF between the tray and the cooling plate. 30. Do not use ddH2 O or tap water! The former leads to very low conductivity between electrode and IPG strip, the latter to very high. 31. Each electrode has a side marked red or black. 32. Program for 24 cm IPG pH 3–10 NL strips using the EPS 3501 XL power supply (adapted from Hoving (40)): Phase 1, 300 V, 1 Vh (0.006 h); Phase 2, 300 V, 900 Vh (3 h); Phase 3, 3500 V, 9500 Vh (5 h), and Phase 4, 3500 V, 52500 Vh (15 h). Current and power are set non-limiting (2 mA, 5 W). Phases 1–4 are programmed in the gradient mode. The voltage will be ramping up to

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Ambort et al. the maximum set in the phase, starting from zero in the first phase and in phases to follow from the end point of the phase before. Therefore in phases 1 and 3 voltage is linearly increased and in phases 2 and 4 held constant. The current check option must be switched off!

Acknowledgments The authors wish to acknowledge and thank Ursula Luginbühl for excellent technical assistance. This work was funded by the Swiss National Science Foundation (SNSF) (grant 3100A0-100772 to E.E.S.) and the European Science Foundation (ESF) Integrated Approaches for Functional Genomics (grant 0341 to D. A.).

References 1. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–021. 2. Klose, J. (1975) Protein mapping by combined isoelectric focusing and electrophoresis in mouse tissues. A novel approach to testing for induced point mutations in mammals. Humangenetik 26, 231–43. 3. Lämmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–85. 4. O’Farrell, P. Z., Goodman, H. M., O’Farrell, P. H. (1970) High-resolution twodimensional electrophoresis of basic as well as acidic proteins. Cell 12, 1133–42. 5. Bjellqvist, B., Ek, K., Righetti, P. G., Gianazza, E., et al. (1982) Isoelectric focusing in immobilized pH gradients: principle, methodology and some applications. J. Biochem. Biophys. Methods 6, 317–39. 6. Görg, A., Postel, W., Günther, S. (1988) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 9, 531–46. 7. Görg, A., Obermaier, C., Boguth, G., et al. (2000) The current state of twodimensional electrophoresis with immobilized pH gradients. Electrophoresis 21, 1037–53. 8. Görg, A., Weiss, W., Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 4, 3665–85. 9. Matsudaira, P. (1987) Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes. J. Biol. Chem. 262, 10035–38. 10. Aebersold, R. H., Leavitt, J., Saavedra, R. A., Hood, L. E., Kent, S. B. (1987) Internal amino acid sequence analysis of proteins separated by one- or twodimensional gel electrophoresis after in situ protease digestion on nitrocellulose. Proc. Natl. Acad. Sci. 84, 6970–74. 11. Rosenfeld, J., Capdevielle, J., Guillemot, J. C., Ferrara, P. (1992) In-gel digestion of proteins for internal sequence analysis after one- or two-dimensional gel electrophoresis. Anal. Biochem. 203, 173–79.

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12. Yates 3rd, J. R., Speicher, S., Griffin, P. R., Hunkapiller, T. (1993) Peptide mass maps: a highly informative approach to protein identification. Anal. Biochem. 214, 397–408. 13. James, P., Quadroni, M., Carafoli, E., Gonnet, G. (1994) Protein identification in DNA databases by peptide mass fingerprinting. Protein Sci. 3, 1347–50. 14. Cottrell, J. S. (1994) Protein identification by peptide mass fingerprinting. Pept. Res. 7, 115–24. 15. Herbert, B. R., Sanchez, J.C., Bini, L. (1997) Two-dimensional electrophoresis: The state of the art and future directions, in Proteome Research: New Frontiers in Functional Genomics (Wilkins, M. R., Williams, K. L., Appel, R. D., Hochstrasser, D. F., eds.), Springer, Berlin, pp. 13–33. 16. Rabilloud, T., Adessi, C., Giraudel, A., Lunardi, J. (1997) Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18, 307–16. 17. Link, A. J. (ed.) (1999) 2-D Proteome Analysis Protocols. Humana, Totowa, NJ. 18. Westermeier, R. (2001) Electrophoresis in Practice, 3rd Edition, Wiley-VCH, Weinheim. 19. Yates 3rd, J. R., Gilchrist, A., Howell, K. E., Bergeron, J. J. (2005) Proteomics of organelles and large cellular structures. Nature 6, 702–14. 20. Lim, J. W. E., Bodnar, A. (2002) Proteome analysis of conditioned medium from mouse embryonic fibroblast feeder layers which support the growth of human embryonic stem cells. Proteomics 2, 1187–1203. 21. Boraldi, F., Bini, L., Liberatory, S., Armini, A., et al. (2003) Normal human dermal fibroblasts: Proteomic analysis of cell layer and culture medium. Electrophoresis 24, 1292–1310. 22. Mears, R., Craven, R. A., Hanrahan, S., Totty, N., et al. (2004) Proteomic analysis of melanoma-derived exosomes by two-dimensional polyacrylamide gel electrophoresis and mass spectrometry. Proteomics 4, 4019–31. 23. Prowse, A. B. J., McQuade, L. R., Bryant, K. J., Van Dyk, D. D., et al. (2005) A proteome analysis of conditioned media from human neonatal fibroblasts used in the maintenance of human embryonic stem cells. Proteomics 5, 978–89. 24. Volmer, M. W., Stühler, K., Zapatka, M., Schöneck, A., et al. (2005) Differential proteome analysis of conditioned media to detect Smad4 regulated secreted biomarkers in colon cancer. Proteomics 5, 2587–2601. 25. Eagle, H. (1959) Amino acid metabolism in mammalian cell cultures. Science 130, 432–7. 26. Richardson, J. C., Scalera, V., Simmons, N. L. (1981) Identification of two strains of MDCK cells which resemble separate nephron tubule segments. Biochim. Biophys. Acta 673, 26–36. 27. Damerval, C., DeVienne, D., Zivy, M., Thiellement, H. (1986) Technical improvements in two-dimensional electrophoresis increase the level of genetic variation detected in wheat-seedling protein. Electrophoresis 7, 53, 54. 28. http://www.millipore.com/userguides.nsf/dda0cb48c91c0fb6852567430063b5d6/6 03b133b9b2a919c85256b3e0050b862/$FILE/P36006.pdf (User guide for Centricon® Plus-70 centrifugal filter devices from Millipore)

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29. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–54. 30. Ramagli, L. S., Rodriguez, L. V. (1985) Quantitation of microgram amounts of protein in two-dimensional polyacrylamide gel electrophoresis sample buffer. Electrophoresis 6, 559–63. 31. Lowry, O. H., Rosebrough, N. J., Farr, A. L., Randall, R. J. (1951) Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–95. 32. Mokrasch, L. C., McGilvery, R. W. (1956) Purification and properties of fructose1, 6-diphosphatase. J. Biol. Chem. 221, 909–17. 33. Smith, R. K., Krohn, R. I., Hermanson, G. T., et al. (1985) Measurement of protein using bicinchoninic adic. Anal. Biochem. 150, 76–85. 34. http://www.piercenet.com/files/1296dh4.pdf (Instructions for BCATM Protein Assay Kit from Pierce) 35. Langen, H., Roder, D., Juranville, J. F., Fountoulakis, M. (1997) Effect of protein application mode and acrylamide concentration on the resolution of protein spots separated by two-dimensional gel electrophoresis. Electrophoresis 18, 2085–90. 36. Rabilloud, T., Valette, C., Lawrence, J. J. (1994) Sample application by ingel rehydration improves the resolution of two-dimensional electrophoresis with immobilized pH gradients in the first dimension. Electrophoresis 15, 1552–58. 37. Sanchez, J. C., Rouge, V., Pisteur, M., Ravier, F., et al. (1997) Improved and simplified in-gel sample application using reswelling of dry immobilized pH gradients. Electrophoresis 18, 324–27. 38. Hoving, S., Voshol, H., van Oostrum, J. (2000) Towards high perfomance twodimensional gel electrophoresis using ultrazoom gels. Electrophoresis 21, 2617–21. 39. Hoving, S., Gerrits, B., Voshol, H., Muller, D., et al. (2002) Preparative twodimensional gel electrophoresis at alkaline pH using narrow range immobilized pH gradients. Proteomics 2, 127–34. 40. http://www1.amershambiosciences.com/applic/upp00738.nsf/vLookupDoc/319 798244-C534/$file/80642960.pdf (Amersham 2-D electrophoresis handbook)

11 Sample Preparation for Mass Spectrometry Analysis of Formalin-Fixed Parafﬁn-Embedded Tissue Proteomic Analysis of Formalin-Fixed Tissue Nicolas A. Stewart and Timothy D. Veenstra

Summary One of the great hopes in biomedical research is that proteomic technology can be used to identify novel biomarkers for diseases such as cancer. The challenge to discovering biomarkers starts with sample collection and continues right through data acquisition and bioinformatic analysis. Because the ultimate goal is to find indicators of human disease it is ideal to be able to study clinical samples. Unfortunately clinical samples such as serum, plasma, urine, and especially tissue biopsies are precious and are often difficult to obtain in sufficient quantities or numbers to conduct proteomic discovery studies. There exists, however, a vast archive of pathologically characterized clinical samples in the form of formalin fixed paraffin embedded tissue blocks. This chapter describes methods that have been developed to allow the proteins from these tissue samples to be excised in a form that is amenable for proteomic analysis by mass spectrometry.

Key Words: Cancer biomarkers; formalin-fixed paraffin embedded tissue; mass spectrometry; proteomics; tissue microdissection.

1. Introduction One of the greatest determinants on the survival rate from any cancer is the stage at which it is detected. The survival rate from cancers detected at an early stage is generally higher but decreases as the stage of detection increases. Therefore a major impetus in proteomics research is to identify biomarkers of early stage diseases. Many of these biomarker discovery efforts are aimed towards the use of biofluids such as serum and plasma. These sample types, From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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however, are inherently difficult to analyze and are often too precious to be given over to proteomic-type discovery studies that are continuing to evolve in both efficacy and success. Another complication is that although biomarkers that originate from the site of a tumor may have a high local concentration, by the time they become diluted within the circulatory system, their effective concentration in the acquired clinical sample may be vanishingly small. Ideally, it would be beneficial to identify a biomarker at the tumor level and determine if it could be translated to a marker detectable in serum or plasma. Tissue or tumor biopsies therefore, are ideal sample candidates for the discovery of potential biomarkers. However, the feasibility of their use for such studies is hampered because of the fact that they are relatively difficult to obtain and require careful storage and handling. Formalin-fixed, paraffin-embedded (FFPE) tissues, on the other hand, represent a vast resource for retrospective protein biomarker investigation. Formalin fixation and paraffin embedding of tissue is practiced in pathology labs worldwide as a standard processing method by which tissues can be stored and catalogued as stable entries. Unfortunately, the storage stability of these tissues partially arises from the high degree of covalently crosslinked proteins. Currently, immunohistochemistry (IHC) is the only published technology capable of providing protein information from these samples (1,2). In addition, because IHC requires a priori knowledge of individual proteins being analyzed, it is not a discovery-based technology. As with tissue biopsies, FFPE sections are heterogeneous in that they contain many different types of cells. To acquire a homogeneous population of cells, laser-capture microdissection (LCM) is used. This technique has the ability to directly isolate a user-defined population of cells from their tissue microenvironment (3). Although LCM of fresh tissue is widely practiced, outside of the interest in conducting tissue microdissection of FFPE tissues for mRNA extraction, microdissection of FFPE tissues is not widely practiced for extraction and analysis of soluble protein (4). Mass spectrometry (MS) is arguably the driving technology in discoverydriven proteomics. Dramatic technical improvements in MS instrumentation combined with the rapid growth of genomic and proteomic databases have enabled development of approaches to identify and quantify large numbers of proteins from complex samples such as serum and tissues (5). Combining LCM of FFPE tissues and MS has the potential for generating protein biomarker data necessary for discovering proteins that are key determinants or indicators of diseases such as cancer. A common misnomer is that MS identifies proteins in proteomic studies. Actually in its present form, MS is best suited to the identification of peptides that are produced from the enzymatic digestion of proteins. It is through

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the analysis of peptide surrogates that proteins are identified. Because of their size, peptides are very amenable to tandem MS (MS/MS) methods that produce partial amino acid sequence information that leads to their identification. Besides the advances made in MS instrumentation, its coupling with nanoflow reversed-phase liquid chromatography (nanoRPLC) has made the analysis of complex peptide mixtures possible. To compare the tryptic peptide abundances obtained from two different FFPE tissues, stable-isotope labeling using trypsin-mediated 16 O/18 O incorporation may also be employed. 2. Materials 2.1. Tissue Processing 1. 2. 3. 4.

Mayer’s hematoxylin stain. Eosin stain. Graded ethanol solutions. SubX organic solvent (Surgipath Medical Industries, Richmond, IL).

2.2. Protein Extraction 1. 2. 3. 4.

50% glycerol. LCM instrument. Low-binding microcentrifuge tubes. The Liquid Tissue™-MS protein prep kit (Expression Pathology, Inc., Gaithersburg, MD).

2.3. Trypsin Digest 1. 2. 3. 4. 5.

Porcine sequencing grade modified trypsin (Promega, Madison, WI). Dithiothreitol (DTT, Sigma, St. Louis, MO). Trifluoroacetic acid (TFA) (≥ 98.0% pure). Incubator for the digest at 37°C in microcentrifuge tubes. High performance liquid chromatography (HPLC)-grade acetonitrile (ACN) (EMD Chemicals Inc., Gibbstown, NJ). 6. C-18 reverse phase microcolumns (e.g. ZipTip®, Millipore, Billerica, MA). 7. Conditioning, loading and eluting buffers for reverse phase desalting microcolumns: ACN, 0.1% TFA in ddH2 O, and 60% ACN, 0.1% TFA solution, respectively.

2.4. LC-MS/MS analysis of tryptic peptides 1. HPLC system capable of NanoRPLC (e.g., Agilent 1100 capillary LC system, Agilent Technologies, Palo Alto, CA). 2. Fused silica capillary column: 75 μm inner diameter × 360 outer diameter × 10 cm long, slurry packed with C-18 silica-bonded stationary phase; 3 μm, 300 Å pore size.

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3. Linear ion trap MS coupled to the NanoRPLC (e.g., LTQ, Thermo Electron, San Jose, CA). 4. High resolution MS coupled to the NanoRPLC for 18 O isotope labeling analysis (e.g., LTQ-FTICR, Thermo Electron, San Jose, CA). 5. Trifluroroacetic acid is diluted to 0.1% (v/v) and is used to resolubilize tryptic peptides before LC-MS/MS. 6. A solution of 0.1 % (v/v) formic acid (FA) in ddH2 O (NANOPure Diamond water system, Barnstead International, Dubuque, IA) (Mobile Phase A) and a solution of 0.1% FA in HPLC-grade ACN (Mobile Phase B) are prepared for the gradient used in the reverse phase liquid chromatographic separation of tryptic peptides.

2.5. Mass Spectrometry Analysis and Bioinformatic Analysis 1. Accompanying software for the collection of the raw MS/MS data generated. 2. Software to search raw data files to a database (e.g., SEQUEST).

3. Methods 3.1. Tissue Processing 1. For tissue microdissection, 10 μm thick tissue sections are cut from FFPE whole mount tissue blocks and placed on coated slides. 2. The section is then heated for 60 min at 58°C. To remove paraffin, treat the section with SubX organic solvent (Surgipath Medical Industries, Richmond, IL) twice for 5 min. 3. The tissue is then rehydrated through multiple, graded ethanol solutions and distilled water. Tissue is then counterstained with Mayer’s hematoxylin, and dehydrated through graded ethanol solutions, and air-dried.

3.2. Laser-capture Microdissection 1. Rehydrate the tissue with 50% glycerol in water for 5 min before laser capture microdissection (LCM). 2. Place the slide containing the tissue upside-down below the objective lens and locate cellular regions with specific histological features of interest (see Note 1). 3. Capture cells with the LCM instrument utilizing an excimer laser (MPB Technologies PSX-100) operating at the following conditions: 248 nm wavelength, 2.5 ns pulse, Emax = 5 mJ, repetition rate = 0.1–100 Hz (see Note 2). 4. Captured cells within the selected area are then transferred into a 1.5 mL low-binding microcentrifuge receiving tube. Approx 100,000–200,000 cells are required for subsequent proteomic analysis (see Note 3).

3.3. Protein Extraction and Trypsin digest 1. Cells collected by microdissection for nano-reversed-phase liquid chromatography-tandem mass spectrometry (RPLC-MS/MS) analysis were

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3. 4. 5. 6. 7.

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processed using proprietary reagents according to the manufacturer’s recommendations (Liquid Tissue™, Expression Pathology Inc., Gaithersburg, MD). The material for nanoRPLC-MS/MS analysis was suspended in 20 μL of Liquid Tissue™ reaction buffer, and incubated for 90 min at 95°C, followed by cooling on ice for 3 min. DTT is added to a final concentration of 10 mM. Heat samples for 5 min at 95°C, and let cool to room temperature. Trypsin (15–18 units) is added and the sample is incubated at 37°C for 18 h. The resulting proteome digestates may be stored at –20°C until analysis. Tryptic peptides generated from the comparative FFPE samples are desalted using C-18 microcolumns. Microcolumns are conditioned by aspirating and dispensing with conditioning buffer (10 μL, 3×), followed by loading buffer (10 μL 3×). Peptide samples are re-solubilized in loading buffer (10 μL) and washed with loading buffer (10 μL 3×). Peptides are eluted from the microcolumn with the elution buffer (10 μL) into microcentrifuge tubes, and lyophilized. Peptide samples are re-solubilized in loading buffer (10 μL) and transferred to vials for autosampler.

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1. For isotope labeling, the protein extracts from equivalent numbers of cells microdissected from FFPE tissues are reconstituted separately in H2 16 O and H2 18 O each containing 20% methanol (v/v). 2. Sequencing grade trypsin is resuspended in the appropriately labeled water (i.e. H2 16 O or H2 18 O), and added to the related sample at an enzyme to protein ratio of 1:20. 3. Incubate at 37°C for 16 h. After this time, an additional equivalent aliquot of the trypsin is added and the samples incubated at 37°C for an additional 6 h (see Note 7). 4. TFA is added to a final concentration of 0.4% (v/v) to stop the reaction (see Note 8). 5. The differentially labeled proteome samples are pooled and lyophilized to dryness. 6. Samples are resolubilized in loading buffer (10 μL) and transferred to vials for autosampler.

3.5. Nanoﬂow RPLC-MS/MS Analysis 1. Nanoflow RPLC is performed on a capillary LC system coupled online to a ion trap MS or an MS capable of high resolution (i.e., 100,000) for quantitation of 16 O/18 O-labeled samples (see Note 9). 2. Wash column for 30 min with 98% mobile phase A at a flow rate of 0.5 μL/min prior to sample injection. 3. Inject sample (typically 1–6 μL) onto the column.

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4. Elute peptides using a linear gradient of 2–40% mobile phase B in 110 min, then to 98% mobile phase B in an additional 30 min, all at a constant flow rate of 0.25 μL/min.

3.6. Mass Spectrometry Analysis and Bioinformatic Analysis 1. The MS is operated in a data dependent MS/MS mode in which each full MS scan is followed by five MS/MS scans of the five most abundant peptide molecular ions. 2. Subject peptides to collision-induced dissociation (CID) using a normalized collision energy of 35% (see Note 10). The heated capillary temperature and electrospray voltage of the mass spectrometer are typically set at 160°C and 1.5 kV, respectively. 3. Although data collection may be dictated based on the available mass to charge (m/z) range of the instrument, it is typically collected over a broad range of 400–2,000. 4. Because the starting amount of protein obtained from the LCM cells is low, multidimensional fractionation prior to MS analysis is inefficient owing to sample handling losses associated with chromatography. 5. To maximize the number of peptides identified, segmented precursor selection scan ranges (i.e., gas phase fractionation in the m/z dimension, GPFm/z ) are used. The following overlapping m/z intervals may be used as a guideline: m/z 400–605, 595–805, 795–1005, 995–1,205, 1,195–1,405, 1,395–1,605, 1,595–1,805, 1,795–2,000, 400–805, and 795–1,200. Data for the 16 O/18 O-labeled experiments are acquired using FTICR detection in centroid mode for the full MS scan (m/z 400–2000) at 100,000 resolution followed by MS/MS of the top five most abundant molecular ions detected. 6. The tandem mass spectra are searched against the UniProt human proteomic database from the European Bioinformatics Institute (http:// www.ebi.ac.uk/integr8) using a combination of database and searching algorithm software (e.g. SEQUEST). Peptides are searched using fully tryptic cleavage constraints and a dynamic 4.008 amu modification on the C-terminus for when 18 O isotope labeling analysis. When using SEQUEST, a legitimately identified peptide should have cross correlation (Xcorr ) scores of 1.9 for [M+H]1+ , 2.2 for [M+2H]2+ and 3.5 for [M+3H]3+ and a minimum delta correlation score (Cn ) of 0.08

4. Notes 1. Microdissection is best performed using software-directed laser pulses to strike at a constant velocity and rate throughput over the previously defined and mapped cellular regions. 2. Target slides are optically transparent quartz coated with an energy transfer coating with the exact dimensions of a standard histology glass slide. The slide stage is a computer-controlled, XY translation stage. A 1/8 beam-splitter is

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4.

5.

6.

7.

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used to split the laser to an energy meter with the remaining beam traveling to an ultraviolet reflective mirror and directed down (-Z) to a ×10 microscope objective (LMU-10X-UVR, OFR). The objective focuses the laser onto the slide and the LCM process is observable using a confocally aligned CCD camera. Because many FFPE tissues have been in storage for years, if not decades, an obvious concern when conducting proteomic investigations of these samples is the effect of long-term storage. Presently there are not enough studies on this topic to make any solid conclusions, however, no obvious detrimental effect has been observed in those studies that have examined FFPE tissues using MS (8,9). One of the studies showed that formalin-fixation and storage does not result in an inordinate degree of oxidation to methionyl and cysteinyl residues nor were any adverse effects on the tryptic digestion efficiency observed (8). Unfortunately the maximum number of cells that can typically be microdissected from FFPE tissues is on the order of 200,000. Although this number will vary depending on the tissue and its heterogeneity, it typically does not yield enough protein to employ multidimensional fractionation before MS analysis. The protein yield for a typical experiment is on the order of 10–20 μg, whereas typically 200 μg is required for a multidimensional fractionation consisting of strong cation exchange prior to RPLC to be employed. There are experimental and data analysis issues related to quantitative proteomics whether it is done using O16 /O18 labeling or subtractively (i.e., when the number of peptides identified from one proteome sample is compared to that identified in another). The trypsin-mediated incorporation of O18 is not always absolutely complete because of the incomplete exchange at the peptide’s carboxy-terminus. Although no absolute reason for this effect has been established, it may be because of the low abundance of the peptide within the complex mixture. Manual analysis of the MS spectrum should always be conducted when a potential interesting abundance change is suggested. When using a subtractive proteomics approach, the precision level is such that abundance changes below three-fold are not considered reliable. A constant issue when dealing with the identification of large numbers of peptides from complex mixtures is the false positive rate. It is impossible to orthogonally validate all of the peptides/proteins identified in these studies, therefore acceptable filtering criteria are necessary when evaluating correct identifications. SEQUEST is a commonly used software program used for identifying peptides based on tandem MS spectra. The filtering criteria based on Xcorr andCn scores are provided as a guideline only. In bioinformatic analyses, they have been shown to provide a false positive rate of approx 1.5% (5). Because the proteins are extracted from the FFPE tissues as tryptic peptides, the role of trypsin during this step is to enzymatically exchange the C-terminal carboxyl oxygen atoms with the appropriate oxygen (i.e., 16 O or 18 O) isotope (6). An alternative means to stop the trypsin-mediated incorporation of 18 O is to boil the sample at the end of the digest. This is acceptable, but the sample should

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not be boiled after addition of acid as deamidation of asparagine or glutamine residues may occur. 9. An MS capable of high resolution (e.g., FTICR-MS) is necessary for accurately quantifying the different isotopically-labeled counterparts from the two proteome samples to be compared. 10. To minimize the selection of peptides that have already been subjected to CID, dynamic exclusion is used. While this is a user defined value, 90 s. is typical.

Acknowledgments This project has been funded in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under Contract N01CO-12400. The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organization imply endorsement by the United States Government. References 1. MacIntyre, N. (2001) Unmasking antigens for immunohistochemistry. Br. J. Biomed. Sci. 58, 190–96. 2. Shi, S. R., Cote, R. J. and Taylor, C. R. (2001) Antigen retrieval techniques: current perspectives. J. Histochem. Cytochem. 49, 931–37. 3. Emmert-Buck, M. R., Bonner, R. F., Smith, P. D., et al. (1996) Laser capture microdissection. Science 274, 998–1001. 4. Gillespie, J. W., Best, C. J., Bichsel, V. E., et al. (2002) Evaluation of non-formalin tissue fixation for molecular profiling. Am. J. Pathol. 160, 449–57. 5. Peng, J. and Gygi, S. P. (2001) Proteomics: the move to mixtures. J. Mass Spectrom. 36, 1083–91. 6. Yao, X., Freas, A., Ramirez, J., Demirev, P. A., and Fenselau, C. (2001) Proteolytic 18O labeling for comparative proteomics: model studies with two serotypes of adenovirus. Anal. Chem. 73, 2836–42. 7. Yates, J. R. III, Eng, J. K., McCormack, A. L., and Schieltz, D. (1995) Method to correlate tandem mass spectra of modified peptides to amino acid sequences in the protein database. Anal. Chem. 67, 1426–36. 8. Hood, B. L., Darfler, M. M., Guiel, T. G., et al. (2005) Proteomic analysis of formalin-fixed prostate cancer tissue. Mol. Cell. Proteomics 4, 1741–53. 9. Crockett, D. K., Lin, Z., Vaughn, C. P., Lim, M. S., Elenitoba-Johnson, K. S. (2005) Identification of proteins from formalin-fixed paraffin-embedded cells by LC-MS/MS. Lab. Invest. 85, 1405–15.

12 Metalloproteomics in the Molecular Study of Cell Physiology and Disease Hermann-Josef Thierse, Stefanie Helm, and Patrick Pankert

Summary Physical and chemical stresses as well as metal-related diseases can disrupt the normal trafficking of metal ions. Moreover, homeostatic imbalance of such metal ions may modulate essential cellular functions (including signal transduction pathways), may catalyze oxidative damage, and may affect the folding of nascent proteins. Here we describe a new qualitative subproteomic method for the detection, isolation, and identification of metal-interacting proteins. Combining both classical immobilized metal ion affinity chromatography (IMAC) and modern proteomic techniques (e.g., two dimensional gel electrophoresis [2-DE]), metal-specific proteins have been successfully isolated and identified to define a metalloproteome. These metal-specific proteomes may give new insights into metal-related pathophysiological processes, such as the allergic reaction to nickel, which represents the most common form of human contact hypersensitivity.

Key Words: 2-DE; 2-dimensional gel electrophoresis; disease proteomics; IMAC; immobilized metal ion affinity chromatography; metal affinity; metalloproteome; nickel; nickel allergy; nitrilotriacetic acid.

1. Introduction Because immobilized metal ion affinity chromatography (IMAC) was first developed for metal-specific protein isolation by Porath et al. (1,2), a large number of IMAC protocols have been developed. All IMAC protocols share the same principal that proteins with exposed histidine and cysteine side chains have a distinct affinity for certain metals, like Ni2+ , Co2+ , Zn2+ , Cu2+ , Fe3+ , or Mn2+ (3,4). Depending on the chosen metal-chelating group and metal ion more or fewer coordination sites are accessible for potential protein binding. From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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In general, binding affinity of proteins to metal ions is dependent on the amino acid composition of a given protein, with histidine showing the highest metal affinity followed by tryptophan, phenylalanine, tyrosine, and cysteine (5). However, the affinity between a metal ion and an amino acid is also highly dependent on the specific metal ion itself, with Fe3+ , for example, having the highest affinity to carboxyl- and phosphate-groups. Besides the amino acid sequence and the distribution of a certain amino acid, surface characteristics and protein folding (3-dimensional structure) are additional important parameters for binding affinity. In an optimal column or bead-based performance the selected metal is strongly held by the matrix bound metal-chelating group, e.g., nitrilotriacetic acid (NTA) or iminodiacetic acid (IDA), still leaving metal coordination sites available to interact with the metal-specific protein ligand. Tetradentate NTA, for example, was found to bind Ni2+ with three carboxyl groups and one nitrogen (6), leaving two other ligand positions accessible to Ni2+ -specific proteins or recombinant 6His-tagged proteins (7). Interestingly, when compared to the dissociation constant (Kd ) of most antibody bindings (Kd from10−5 M to 10−12 M), 6His-tagged protein to Ni2+ Kd has been shown to be relatively high, 10−13 M at pH 8 (8,9). Moreover, Ni-NTA itself has been shown to be stable over a pH range of 2.5–13 and withstands extreme conditions like 2% SDS and 100% ethanol. Today, IMAC is not only commonly used for purification of the mentioned recombinant 6His-tagged proteins or the evaluation of protein folding and endotoxin removal from protein preparations, but also advantageous in the enrichment of phosphopeptides (10). Reversible protein phosphorylation (e.g., at Ser, Thr, and Tyr residues) is one of the most important post-translational modifications, controlling signal transduction pathways that direct cellular activation, differentiation, and proliferation, as well as apoptosis (11). Because phosphopeptides are acidic and show strong binding characteristics to some metal ions (Fe3+ , Ga3+ , and Al3+ ) different IMAC protocols have been introduced for isolation of phosphopeptides (10,12). Among the most recently described phosphopeptide IMAC methods are titanium dioxide (TiO2 ) phosphopeptide enrichment (13,14), and metal oxide affinity chromatography (MOAC) where the affinity of the phosphate group for Al(OH)3 is exploited (15). For detailed information on IMAC phosphopeptide enrichment see Corthals et al., 2005 and Ueda et al., 2003 (5,10). Affecting up to 15% of the women in industrialized countries, human nickel (Ni) allergy represents one of the major metal-related diseases in the human population. However, according to several studies transition metal Ni is two-faced. Nickel is considered as a beneficial physiological agent used for essential functions as an ultra trace element. Conversely, Nickel can act as a pathological agent by interacting directly with DNA or DNA-binding proteins

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or metal-specific T cells, thus causing cellular toxicity or a metal-specific allergic contact dermatitis (ACD). To elucidate such physiological and diseaserelated molecular processes, a new qualitative subproteomic method, combining IMAC and 2-dimensional gel electrophoresis (2-DE), has been developed to enrich and identify Ni2+ -interacting proteins from different human cell types (e.g., human antigen presenting cells, keratinocytes). After affinity binding of metal-interacting proteins to Ni-NTA beads, followed by stringent washing steps and elution with imidazole, proteins are applied to modern 2-DE using immobilized pH-gradients (16–19). In a typical proteomic workflow, proteins are identified by mass spectrometry to define the metalloproteome. Complementary methods such as the use of recombinant proteins and metal detection by atomic absorption spectrometry allow confirmation of direct metal-protein interactions (20). Thus, with a clear differential pattern to Cu-binding proteins in human liver cells, Ni-NTA affinity enrichment of proteins from human B-cell lysates resulted in the subproteomic identification of so far unknown Ni-interacting proteins (Fig. 1) (21–23). Quite unexpectedly, 16 of these Niinteracting proteins were found to belong to the group of stress-inducible heatshock proteins or chaperonins, including HSP-60, HSP-70, BIP, and the high-molecular heterooligomeric complex of TriC/CCT (21). Thus Ni2+ , in addition to the induced formation of T cell epitopes recognizable by the A

B Protein solution cell lysate 0.1% Triton X-100

Ni-NTA-beads

pH 4

pH 7

MW

1h incubation, 4°C

Washing steps 500 mM NaCl (high salt) 137 mM NaCl (low salt)

Elution of Ni-interacting proteins 250 mM imidazole

2-DE, staining, spot picking, trypsinization

Fig. 1. Ni-affinity enriched proteins were isolated from human antigen presenting cells (2* 107 cells) as demonstrated in the subproteomic workflow (A) and analyzed by 2-DE (B, silver staining), and subsequently identified by mass spectrometry (for details of mass spectrometric protein identification see (21)).

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acquired immune system, intimately interacts with essential constituents of the innate defense system, thereby linking both arms of the immune system. In summary, the development and combined usage of new metal-specific and proteomic techniques gives new insights into metal-related metabolic pathways and frequent metal-specific disorders, like human nickel ACD, leading to improved strategies in diagnosis and therapy of such environment-induced disease (21,24,25). 2. Materials 2.1. Cell Culture and Lysis 1. RPMI 1640 medium for human antigen presenting cells containing 10% fetal calf serum, 2 mM l-glutamine, 1 mM sodium pyruvate, nonessential amino acids and 25 mM HEPES buffer (all from Gibco BRL Life Technologies, Paisley, UK). For primary keratinocytes Keratinocyte Basal Medium 2 was supplemented with SupplementPack/Keratinocyte Growth Medium 2 (KGM) (Promocell, Heidelberg, Germany). 2. A solution of trypsin (0.1%) and ethylenediamine tetraacetic acid (EDTA) (0.02%) (Biochrom, Berlin, Germany). 3. Phosphate buffered saline (D-PBS) (Invitrogen, Karlsruhe, Germany). 4. Fetal calf serum (FCS) (Biochrom, Berlin, Germany). 5. Lysis buffer (0.1% Triton): 137 mM NaCl, 20 mM Tris-HCl, 10% (v/v) glycerol, 0.1% (v/v) Triton X-100, pH 8.2. Add protease inhibitor cocktail tablets, “complete mini, EDTA free” (Roche, Mannheim, Germany) before use. Store in aliquots at –20°C (see Note 1).

2.2. Isolation of Nickel-Binding Proteins 1. Nickel-nitrilotriacetic acid (Ni-NTA) Magnetic Agarose Beads 5% suspension, with binding capacity: 300 μg/mL (Qiagen, Hilden, Germany). 2. High salt lysis buffer: 500 mM NaCl, 20 mM Tris-HCl, pH 8.2, 10% (v/v) Glycerol, 0.1% Triton X-100. 3. Low salt lysis buffer: 137 mM NaCl, 20 mM Tris-HCl, pH 8.2, 10% (v/v) Glycerol, 0.1% Triton X-100. 4. Imidazole solution: 250 mM imidazole (Sigma, Taufkirchen, Germany) in distilled water. 5. Magnetic device for 1.5 mL Eppendorf tubes (Qiagen, Hilden, Germany). 6. Standard 2-D electrophoresis equipment.

2.3. Silver Staining of Metal-Afﬁnity Enriched Proteins Compatible for Mass Spectrometry (MS) Several protocols of 2-dimensional gel electrophoresis (2-DE) are useful for detection of IMAC separated proteins, without negatively affecting the

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principal method described here (16,17,21). Nevertheless, after separating affinity-enriched proteins by 2-DE a protein staining method compatible for mass spectrometric analysis has to be applied. Following protein gel scanning, e.g., with the LabScan Image Scanner (GE Healthcare, München, Germany) or the new laser scanner FLA 5100 (FUJIFILM Life Science, Düsseldorf), spot picking can be performed by hand or automatically using the PROTEINEER spII system (Bruker Daltonics, Bremen, Germany). Subsequent MALDI mass spectra can be recorded e.g., with a Ultraflex MALDI-TOF spectrometer (Bruker Daltronics, Bremen, Germany) equipped with a 337 nm nitrogen laser (for details see (21)). 1. Thiosulfate solution: 0.02% (w/v) sodium thiosulfate pentahydrate (Merck, Darmstadt, Germany). 2. Silver staining solution: 0.2% (w/v) silver nitrate (Merck, Darmstadt, Germany), 0.02% formaldehyde solution (34%) (J.T. Baker, Deventer, NL). 3. Developer: 3% (w/v) sodium carbonate (J.T. Baker, Deventer, NL), 0.05% formaldehyde solution (37%) (J.T. Baker, Deventer, NL), 0.0005% thiosulfate solution. 4. Stopping solution (suitable for mass spectrometry) (see Note 2): 50% methanol (Merck, Darmstadt, Germany), 12% acetic acid (glacial) (Merck, Darmstadt, Germany). 5. Stopping solution (not suitable for mass spectrometry): 0.5% glycine (Roth, Karlsruhe, Germany). 6. Ethanol.

3. Methods 3.1. Preparing Samples for Isolation of Nickel-Binding Proteins 1. Culture of immune cells in RPMI medium and/or primary keratinocytes in KGM in 10 mm tissue culture dish until passage 3 or earlier. 2. When e.g., keratinocytes are confluent, wash with PBS and incubate with 3 mL of trypsin/EDTA for 5 min at 37°C. Rinse cells with a pasteur pipet and transfer into a tube supplied with 6 mL 10% FCS in PBS. Rinse the dish with additional 3 mL of 10% FCS and transfer it into the tube. 3. Wash cells 3× with cold PBS and count the cells. 4. Resuspend cells in lysis buffer (2* 107 /mL), incubate at 4°C for 1 h with gentle shaking. 5. Clarify the lysate by centrifugation (20,000g, 10 min, 4°C). (see Note 3) 6. The supernatant can be stored in aliquots at –20°C or –80°C.

3.2. Isolation of Nickel-Binding Proteins 1. Resuspend the Ni-NTA Magnetic Agarose Beads by vortexing or pipeting. 2. Incubate 150 μL Ni-NTA Beads with 1 mL lysate (recommended ratio) by rotation for 2 h at 4°C in a 1.5-mL Eppendorf tube (see Note 4).

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3. Insert the tube into a magnetic device for 1.5-mL Eppendorf tubes and remove supernatant (see Note 5). 4. Wash pellet 2× with high salt lysis buffer, once with low salt lysis buffer. The pellet has to be resuspended thoroughly during each wash. Discard supernatants (see Note 6). 5. For elution add the imidazole solution to the pellet and incubate for 10 min at room temperature. Use a volume equal to the original volume of Ni-NTA beads. 6. Insert the tube into a magnetic device for 1.5-mL Eppendorf tubes and carefully transfer the eluate into a fresh tube. Store aliquots at –80°C. 7. For first-dimension isoelectric focussing sample (e.g., 25 μg) can be applied by including it in the rehydration solution of the IPG strip, followed by standard protocols of 2-D electrophoresis.

3.3. Silver Staining of Metal-Afﬁnity Enriched Proteins Compatible for Mass Spectrometry (MS) After 2-DE of metal-affinity enriched proteins spots have to be stained with staining protocols adapted to mass spectrometric analysis (for details see (21)). Silver staining has to be performed in glass dishes and see also Notes 7 – 11. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Fix the gel in 40% ethanol/10% (v/v) acetic acid for at least 1 h. Wash 3× in 30% (v/v) ethanol for 20 min. Reduce the gel in thiosulfate solution for 1 min. Wash 3× in distilled water for 20 sec. Stain the gel in staining solution for 20 min. Wash 3× in distilled water for 20 sec. Develop in developer for 3–5 min. Wash 3x in distilled water for 30 sec. Stop the reaction in stopping solution for 5 min. Wash 2× in distilled water for 30 sec and an additional time for 15 min. Gel can be stored in 1% (v/v) acetic acid or shrink-wrap. An example result is demonstrated in Fig. 1.

Alternatively, gels (e.g., loaded with 100 μg protein concentration) may be stained by MS-compatible Coomassie staining, e.g., according to Jungblut et al. (26). 4. Notes 1. Freshly prepared lysis buffer can be stored at 4°C for several months under sterile conditions. 2. The color of the gel deepens after a while. 3. At this point a determination of the protein concentration is often useful. 4. Incubation of the cell-lysate with Ni-NTA Magnatic Agarose Beads can be extended to overnight incubation.

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5. The supernatant contains all proteins that do not bind to Nickel and maybe additional Nickel-binding Proteins. Therefore, store the supernatant at –20°C or –80°C just in case. 6. Supernatants of the washing steps can be stored at –20°C or –80°C just in case. 7. All steps should be performed at room temperature with gentle shaking. 8. Because the silver stain is very sensitive, nitrile gloves are recommended. Latex gloves may leave fingerprints on the gel. 9. The amount of fluid per gel depends on the size of the gel and the staining dishes. Approximately 250 mL for 20 × 20 cm gels is sufficient. 10. The gel becomes less flexible through the staining procedure and tears very easily. To transport the gel safely onto the scanning device, very careful handling is required. 11. All solutions, containing methanol or silver nitrate have to be disposed of according to the directions given by the local authorities.

Acknowledgments We thank Doris Wild and Stefanie Eikelmeier for excellent technical assistance, and Dr. Ian Haidl, Depts. of Pediatrics, Microbiology and Immunology, Halifax, Canada, for very careful reading of the manuscript. This work was supported in part by the Landesstiftung Baden-Wüerttemberg, Germany, Forschungsprogramm “Allergologie” by grant P-LS-AL/26 (to HJT), and the European Union, as part of the project Novel Testing Strategies for In Vitro Assessment of Allergens (Sens-it-iv), LSHB-CT-2005 – 018681, (www.sensit-iv.eu).

References 1. Porath, J., Carlsson, J., Olsson, I., and Belfrage, G. (1975) Metal chelate affinity chromatography, a new approach to protein fractionation. Nature. 258, 598–9. 2. Porath, J. (1992) Immobilized metal ion affinity chromatography. Protein Expr. Purif. 3, 263–81. 3. Mondal, K. and Gupta, M. N. (2006) The affinity concept in bioseparation: evolving paradigms and expanding range of applications. Biomol. Eng. 23, 59–76. 4. Sun, X., Chiu, J. F., and He, Q. Y. (2005) Application of immobilized metal affinity chromatography in proteomics. Expert Rev. Proteomics. 2, 649–57. 5. Ueda, E. K., Gout, P. W., and Morganti, L. (2003) Current and prospective applications of metal ion-protein binding. J. Chromatogr. A. 988, 1–23. 6. Hochuli, E., Dobeli, H., and Schacher, A. (1987) New metal chelate adsorbent selective for proteins and peptides containing neighbouring histidine residues. J. Chromatogr. 411, 177–84. 7. Terpe, K. (2003) Overview of tag protein fusions: from molecular and biochemical fundamentals to commercial systems. Appl. Microbiol. Biotechnol. 60, 523–33.

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8. Harlow, E., and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. 9. Schmitt, J., Hess, H., and Stunnenberg, H. G. (1993) Affinity purification of histidine-tagged proteins. Mol. Biol. Rep. 18, 223–30. 10. Corthals, G. L., Aebersold, R., and Goodlett, D. R. (2005) Identification of phosphorylation sites using microimmobilized metal affinity chromatography. Methods Enzymol. 405, 66–81. 11. Bollen, M. and Beullens, M. (2002) Signaling by protein phosphatases in the nucleus. Trends Cell Biol. 12, 138–45. 12. Zhou, W., Merrick, B. A., Khaledi, M. G., and Tomer, K. B. (2000) Detection and sequencing of phosphopeptides affinity bound to immobilized metal ion beads by matrix-assisted laser desorption/ionization mass spectrometry. J. Am. Soc. Mass Spectrom. 11, 273–82. 13. Hata, K., Morisaka, H., Hara, K., et al. (2006) Two-dimensional HPLC on-line analysis of phosphopeptides using titania and monolithic columns. Anal. Biochem. 350, 292–7. 14. Larsen, M. R., Thingholm, T. E., Jensen, O. N., Roepstorff, P., and Jorgensen, T. J. (2005) Highly selective enrichment of phosphorylated peptides from peptide mixtures using titanium dioxide microcolumns. Mol. Cell Proteomics. 4, 873–86. 15. Wolschin, F., Wienkoop, S., and Weckwerth, W. (2005) Enrichment of phosphorylated proteins and peptides from complex mixtures using metal oxide/hydroxide affinity chromatography (MOAC). Proteomics. 5, 4389–97. 16. Gorg, A., Obermaier, C., Boguth, G., et al. (2000) The current state of twodimensional electrophoresis with immobilized pH gradients. Electrophoresis. 21, 1037–53. 17. Gorg, A., Weiss, W., and Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics. 4, 3665–85. 18. Klose, J. (1975) Protein mapping by combined isoelectric focusing and electrophoresis of mouse tissues. A novel approach to testing for induced point mutations in mammals. Humangenetik. 26, 231–43. 19. O’Farrell, P. H. (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–21. 20. Thierse, H. J., Moulon, C., Allespach, Y., et al. (2004) Metal-protein complexmediated transport and delivery of Ni2+ to TCR/MHC contact sites in nickelspecific human T cell activation. J. Immunol. 172, 1926–34. 21. Heiss, K., Junkes, C., Guerreiro, N., et al. (2005) Subproteomic analysis of metalinteracting proteins in human B cells. Proteomics. 5, 3614–22. 22. She, Y. M., Narindrasorasak, S., Yang, S., Spitale, N., Roberts, E. A., and Sarkar, B. (2003) Identification of metal-binding proteins in human hepatoma lines by immobilized metal affinity chromatography and mass spectrometry. Mol. Cell. Proteomics. 2, 1306–18. 23. Smith, S. D., She, Y. M., Roberts, E. A., and Sarkar, B. (2004) Using immobilized metal affinity chromatography, two-dimensional electrophoresis and mass spectrometry to identify hepatocellular proteins with copper-binding ability. J. Proteome Res. 3, 834–40.

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24. Kulkarni, P. P., She, Y. M., Smith, S. D., Roberts, E. A., and Sarkar, B. (2006) Proteomics of metal transport and metal-associated diseases. Chemistry. 12, 2410–22. 25. Martin, S. F., Merfort, I., and Thierse, H. J. (2006) Interactions of chemicals and metal ions with proteins and role for immune responses. Mini Rev. Med. Chem. 6, 247–55. 26. Jungblut, P., Baumeister, H., and Klose, J. (1993) Classification of mouse liver proteins by immobilized metal affinity chromatography and two-dimensional electrophoresis. Electrophoresis. 14, 638–43.

13 Protein Extraction from Green Plant Tissue Ragnar Flengsrud

Summary A method for preparation of protein from green plant tissue for two-dimensional electrophoresis is described. The method is demonstrated on barley leaves, potato leaves and spruce needles and appears to overcome the obstacles inherent in green plants to proteomic analysis. The yield and the representation of proteins are discussed.

Key Words: Barley leaves; green plant tissue; potato leaves; protein extraction; spruce needles; two-dimensional electrophoresis.

1. Introduction Preparation of proteins for two-dimensional (2-D) electrophoresis is an important, and sometimes crucial, part of this central proteomics technique, a fact that may be overlooked or underestimated. Plants and especially green plant tissues constitute considerable challenges here, because of low protein concentration and the presence of deleterious compounds in the cell. Plant proteases may contribute to the challenge because remarkable stability has been reported relevant to temperature (1–3), pH (2,3) and even urea or guanidine hydrochloride (2,4). During the work with green plant tissues several principles were applied that seem to overcome these problems, resulting in good, reproducible 2-D separation of green tissue proteins from three different species (5). These principles are: (a) Cell disruption and homogenization in liquid N2 and insoluble polyvinylpyrrolidone; (b) an extraction buffer including thiourea and SDS with a pH lower than 5.5; (c) protein precipitation in 90% (v/v) acetone at –20°C; From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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(d) dialysis against a buffer containing 9.5M urea, nonionic detergent and lysine. The sample preparation method described in this chapter is well suited for barley leaves (Fig. 1), potato leaves and spruce needles. Electropherograms, both in the isoelectric focusing (Fig. 1) and the nonequilibrium pH gradient mode, is presented. There is no indication of protease activity. Total recovery of protein is 6.7–16.5% (5). Yet, the possibility exists that all or most proteins are represented in the extract. A study on the effect of different extraction solutions on the solubilization of endosperm proteins showed that the same proteins were extracted, but to different degree. SDS/urea was one of the extraction solutions with best overall results in this work. Even with 22% protein recovery, the extract was shown to contain hordein proteins (6). The sample preparation method was utilized in a stress diagnosis study on spruce needles (7,8). This study used both soluble and immobilized pH gradients and about 1,500 spots were detected by image analysis. Here, the changes in needle protein pattern were studied by image analysis of 300–350 spots. Protease activity was found to be negligible in this study.

Fig. 1. Two-dimensional electrophoresis in the isoelectric focusing mode of proteins extracted from leaves of a mutant line (H354-33-7-5). of the barley variety cv. Carlsberg II. The loading was 33 μg protein and silver staining was used for detection.

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2. Materials 1. Extraction buffer: 50 mM pyridine, 10 mM thiourea, 1% (w/v) SDS, adjusted to pH to 5.0 by HCl. The purity of pyridine (see Note 1), the pH-range (see Note 2) and the use of SDS (see Note 3) is important. 2. Lysis solution: 9.5M urea, 2% (v/v) nonionic detergent (Igepal CA-630), 1.6% (v/v) ampholytes pH 5–7, 0.4% ampholytes pH 3–10, 2.5% (w/v) dithiothreitol (DTT). 3. Modified lysis solution for dialysis: 2% (w/v) lysine substitutes the ampholytes.

3. Methods 1. Cut the leaves into 0.5 to 1.0 cm parts and needles into 2–4 mm parts. Typically, start with 0.4 g green tissue and grind it twice in a mortar with liquid N2 to give a fine powder. The storage of plant material before homogenization should be considered (see Note 4). 2. Add an amount of insoluble polyvinylpyrrolidone (Polyclar AT) twice the weight of plant material (see Note 5) and mix. 3. Add extraction buffer (9.5 mL) to the mixture in the mortar, stir for a few minutes and centrifuge at +5°C for 40min. at 8,000g. 4. Transfer the supernatant to a thick-walled glass centrifuge-tube, add ice cold acetone (see Note 6) to give a final concentration of 90% (v/v) and mix well. Allow proteins to precipitate at –20°C for 2 h. The yield of the protein depends on the concentration of acetone (see Note 7). 5. Collect proteins by centrifugation for 20 min at +5°C and 5,000g. Discard the supernatant and wash the precipitate once with ice cold acetone and centrifuge as above. 6. Carefully dry the resulting precipitate in a stream of N2 , add 400 μL of the lysis solution and mix well. 7. Dialyse the mixture of precipitate and lysis solution overnight against 25 mL of the modified lysis solution. 8. Centrifuge the dialysed sample at 8,000g for 10 min. Add the appropriate ampholytes to the clear supernatant to give a final 2% (v/v) concentration and store at –20°C in suitable aliquots. The suitability depend on the detection method to be used following the 2-D electrophoresis (see Note 8).

4. Notes 1. The pyridine in the extraction buffer should be distilled over ninhydrin and stored under nitrogen. Alternatively, at least HPLC-grade is used and stored under nitrogen. 2. The study of thiourea as phenoloxidase inhibitor concluded that the pH in the extraction buffer should not be above 5.5 (9). 3. Extraction with and without SDS in the extraction buffer showed that SDS was necessary for the solubilisation of membrane-bound proteins (5).

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4. Storage of barley leaves at –20°C or –80°C up to 3 mo and at +5°C for 1 wk, before homogenization does not seem to affect the protein pattern. 5. The original study (5) used twice the amount (g/g) of insoluble polyvinylpyrrolidone (Polyclar AT) to plant tissue for its homogenization. A later work (7) used routinely equal amounts and observed that less polyvinylpyrrolidone resulted in poor electropherograms. 6. Acetone is kept at –20°C before its use in protein precipitation. 7. A final concentration of 90% acetone at –20°C for 1 h will totally precipitate the proteins (10). 8. The following aliquots of the extract are suitable for 2-D electrophoresis: for silver staining, 40–60 μL, corresponding to 30–46 μg protein; for Coomassie Blue staining, 150–260 μL. The aliquots should not be refrozen.

References 1. Fahmy, A. S., Ali, A. A., and Mohammed, S. A. (2004) Characterization of a cysteine protease from wheat Triticum aestivum (cv. Giza 164). Bioresour. Technol. 91, 297–304. 2. Kaneda, M., Yonezawa, H., and Uchikoba, T. (1995) Improved isolation, stability and substrate specificity of cucumisin, a plant serine endopeptidase. Biotechnol. Appl. Biochem. 22, 215–22. 3. Patel, B. K. and Jagannadham, M. V. (2003) A high cysteine containing thiol proteinase from the latex of Ervatamia heyneana: purification and comparison with ervatamin B and C from Ervatamia coronaria. J. Agric. Food Chem. 51, 6326–34. 4. Uchikoba, T., Niidome, T., Sata, I., and Kaneda, M. (1993) Protease D from the sarcocarp of honeydew melon fruit. Phytochemistry 33, 1005–08. 5. Flengsrud, R. and Kobro, G. (1989) A method for two-dimensional electrophoresis of proteins from green plant tissues. Anal. Biochem. 177, 33–6. 6. Flengsrud, R. (1993) Separation of acidic endosperm proteins by two-dimensional electrophoresis. Electrophoresis 14, 1060–66. 7. Davidsen, N. B. (1995) Two-dimensional electrophoresis of acidic proteins isolated from ozone-stressed Norway spruce needles (Picea abies L. Karst): Separation method and image prosessing. Electrophoresis 16, 1305–11. 8. Davidsen, N. B. (1996) Improved two-dimensional electrophoretic separation of acidic proteins extracted from Norway spruce needles by using immobilized pH gradients. Electrophoresis 17, 1280–81. 9. Van Driessche, E., Beeckmans, S., Dejaegere. R., and Kanarek, L. (1984) Thiourea: the antioxidant of choice for the purification of proteins from phenol-rich plant tissues. Anal. Biochem. 141, 184–88. 10. Neuhoff, V. (1973) Micromethods in Molecular Biology. Springer-Verlag (Kleinzeller, A., Springer, G.F., and Wittmann, H.G., eds.) 14, p133.

14 The Terminator: A Device for High-Throughput Extraction of Plant Material B. M. van den Berg

Summary The Terminator is a device for cost-efficient high-throughput homogenization of plant material and sample preparation. Protein and DNA samples can easily be prepared from large numbers of crude material for further analysis such as protein electrophoresis or polymerase chain reaction (PCR) followed by DNA electrophoresis. The functioning of the device is based on vibration of 96 stainless steel pegs in wells of a standard 96-well micro plate. Using the Terminator all types of plant tissue, including seeds, can be homogenized in standard micro plates in 3 min.

Key Words: DNA extraction; electrophoresis; high throughput analysis; plant material; protein extraction; seeds.

1. Introduction High-throughput homogenization and sample preparation is often a timelimiting step in large-scale analysis of biological material. This certainly holds for the analysis of seed or other plant tissue in several agricultural businesses— for example the seed business (1)—where many hundreds or even thousands of individual samples are daily analyzed on a routine basis. In the seed business, gel isoelectric focusing of protein (but also other electrophoretic techniques) is widely used to determine the genetic purity of seed lots or the percentage of inbreds that may occur in hybrid seed lots. With the advance of isoelectric focusing in the 1980s, the need for high-throughput sample preparation techniques and equipment became important (1). During the From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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years 1988 and 1989 a device was developed—later named the Terminator— that is perfectly suited to address the needs of high-throughput sample preparation for isoelectric focusing of protein (2). In more recent years, the Terminator has proven to be an efficient tool also in high-throughput extraction of DNA from seed and other plant tissue (1,3). In 2004, the Terminator was improved. The device was restyled, another power supply was added, and the design of the stainless steel pegs was improved to increase the efficiency of homogenization. Here, the functioning of the Terminator is presented in detail. 2. Materials All chemicals were purchased from Sigma (St. Louis, OH, USA), unless otherwise stated. 2.1. Sample Preparation 1. The Terminator used is the restyled and improved Terminator as produced and sold by Elexa (Enkhuizen, the Netherlands). This device consists of three parts: the Terminator base plate (Fig. 1A), the Terminator head consisting of a vibromotor and an aluminum plate with 96 stainless steel pegs (Fig. 1B), and a variable power supply (Fig. 1C). The base plate is a 4 cm thick stainless steel plate equipped with a micro plate holder consisting of an aluminum base, plastic holders, and micro plate clips. Standard 96-wells micro plates can be placed on the micro plate holder and fixed with the special clips. The heavy weight of the steel plate and the rubber feet assure that the Terminator remains at a fixed position during operation.

Fig. 1. Image showing the three parts of the Terminator. A, the Terminator base plate; B, the Terminator head consisting of the vibrating motor and the 96-peg plate; C, the variable AC power supply (0–220 V).

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2. 3. 4.

5. 6. 7.

8.

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The Terminator head consists of a special design vibromotor that operates at a vibration frequency of 50 Hz, and an aluminum plate attached to it which bears 96 stainless steel pegs that fit perfectly in the wells of a standard micro plate. The variable power supply is a standard AC 220 V transformer that can deliver 0–220 V (see Notes 1–3). 96-well flat-bottom micro plates (see Note 4) from Costar (Cambridge, USA). Seeds from Seminis Vegetable Seeds (Oxnard, CA, USA) and Syngenta (Nerac, France). Sunflower and corn seed protein extraction buffer for isoelectric focusing (IEF): 10 mM Tris-HCl, pH 7.0, 2% (v/v) ampholytes of same pH range as that of the gel. Make fresh just before use. Tomato seed ADH (alcohol dehydrogenase) extraction buffer: 2% ampholytes (v/v) pH range 3–10, 0.25% (w/v) dithiothreitol. Make fresh just before use. Ampholytes: SinuLytes 3-7 and 3-10 from Sinus (Heidelberg, Germany). Store at 4°C (see Note 5). DNA seed extraction buffer (XB): 0.2M Tris-HCl pH 8.0, 0.5% (w/v) sodium dodecyl sulphate (SDS), 0.3M NaCl, 25 mM ethylenediamine tetraacetic acid (EDTA). Store at room temperature. Protein precipitation (PP) buffer: 2.5M potassium acetate pH 6.5. Add 245.0 g potassium acetate to 800 mL water. Stir until fully dissolved. Adjust the pH to 6.5 with acetic acid. Bring the final volume to 1 L. Store at room temperature. Tris-EDTA (TE) buffer: dissolve 1.21 g Tris and 37.2 mg EDTA in 1 L water. Store at 4°C.

2.2. Isoelectric Focusing (IEF) These instructions assume knowledge of making thin horizontal isoelectric focusing gels between glass plates and the use of equipment for horizontal isoelectric focusing (4–7). 1. 2. 3. 4. 5. 6. 7. 8. 9.

10.

16% (w/v) glycerol. Store at 4°C. 30% acrylamide/bis-acrylamide solution (29:1). Store at 4°C. SinuLytes 3-7 and 3-10 (Sinus, Heidelberg, Germany). Store at 4°C. 10% (w/v) ammonium persulfate. Store at 4°C, but no longer than 1 wk. Electrode paper: Whatman #17 Electrode solution: 2% (v/v) ampholytes with pH-range identical to that of the gel. 96-well sample applicator strips from Elexa (Enkhuizen, The Netherlands). Gel backing: GelGrip from Sinus (Heidelberg, Germany). Coomassie gel staining solution: 0.2% (w/v) coomassie brilliant blue (CBB), 50% (v/v) water, 40% (v/v) ethanol, 10% (v/v) acetic acid. Add 0.1 g coomassie brilliant blue to 20 mL ethanol. Stir until fully dissolved. Then add 25 mL water and 5 mL acetic acid. Prepare fresh before use. CBB destaining solution: 50% (v/v) water, 40% (v/v) ethanol, 10% (v/v) acetic acid. Store at room temperature.

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11. ADH staining solution: 0.1M Tris-HCl, pH 7.5, 5% (v/v) ethanol, 0.2% -nicotinamide adenine dinucleotide (NAD), 0.2% (w/v) 1-(4,5-dimethylthiazol2-yl)-3,5-diphenylformazan (MTT), 0.05% (w/v) phenazine methosulfate (PMS). Prepare fresh. 12. 20% (w/v) trichloroacetic acid (TCA). Prepare fresh.

2.3. Inter Simple Sequence Repeat PCR (ISSR-PCR) ISSR-PCR is a technique based on amplification of DNA between two simple sequence repeat (SSR; head-to-tail tandem arrays of short DNA repeat motifs) regions, that uses 5 or 3 anchored SSR PCR primers (8). 1. Anchored primer: 5’-DVDTCTCTCTCTCTCTC (D = A,G,T ;V = A,C,G) from Invitrogen (Carlsbad, CA, USA). 2. PCR mix: 9.06 μL water, 1.50 μL PCR buffer (10x), 0.60 μL 50 mM magnesium chloride, 0.6 μL dNTP mixture (2.5 mM each), 0.12 μL 25 μM primer, 0.12 μL DNA polymerase, 3 μL pepper DNA solution. The PCR buffer (10×) and the magnesium chloride (MgCl2 ) come at the appropriate concentration with the DNA polymerase. The dNTP’s and primer must be diluted to the appropriate concentration with water. 3. DNA polymerase from Invitrogen (Carlsbad, CA, USA). 4. PCR reaction plates from Invitrogen (Carlsbad, CA, USA).

2.4. DNA Electrophoresis These instructions assume knowledge of making thin horizontal gels between glass plates and the use of equipment for horizontal electrophoresis (3). 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Gel buffer (2×): 120 mM Tris-formic acid, pH 9.0. 10% (w/v) ammonium persulfate. Store at 4ºC, but no longer than 1 wk. 20% (w/v) glycerol. Store at 4ºC. 30% acrylamide/bis-acrylmide solution (29:1). Store at 4ºC. Gel backing: GelGrip from Sinus (Heidelberg, Germany). Sample buffer for DNA electrophoresis (SB): 2.5% gel buffer (2×), 0.02% Bromophenol Blue. In-gel well template and spacers (0.2 mm thick) from Elexa (Enkhuizen, The Netherlands). Electrode paper: Whatman #17. Electrode solution: 10% (w/v) Tris-Base, 1.73% (w/v) boric Acid, 0.02% bromophenol blue. Fixing solution: 2% (v/v) nitric acid. Staining solution: 0.2% (w/v) silver nitrate. Developer solution: 0.05% (v/v) formaldehyde, 3% (w/v) sodium carbonate. Prepare fresh just before use. Stop solution: 5% (v/v) acetic acid.

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3. Methods 3.1. Sample Preparation The Terminator homogenizes tissue in the wells of the micro plate by the vibration in all directions of the pegs in the wells. Fig. 2A illustrates a diagrammatic view of how plant tissue is squeezed and homogenized between the vibrating pegs and the wall and bottom of the micro plate wells. Optimal voltage for operation of the Terminator lies between 90 and 130 V depending on the type of tissue to be homogenized. For locations with 110 net Voltage, a pretransformer that gives 220V output can best be used. 1. Small parts of the corn and sunflower seeds are punched using a home-made device to get samples that fit in the wells of a 96-well plate (see Note 7). Alternatively, parts can be cut using a scalpel. To cut the seed parts easier, the seeds may be soaked in water overnight. The seed parts are put in the wells of a 96-well micro plate, 200 μL seed extraction buffer is added, and the tissue is disrupted and fully homogenized using the Terminator. After 3 min operation of the Terminator the homogenates are centrifuged in a micro plate centrifuge and the supernatant samples are used for IEF. 2. Tomato seeds are put in the wells of a 96-well micro plate, 200 μL extraction buffer is added, and the seeds are disrupted and fully homogenized using the Terminator.

Fig. 2. Composite image showing a close view of the pegs of the Terminator in micro plate wells. A drawing of a peg in a well of a micro plate illustrating how tissue is squeezed between the pegs and the walls and bottom of the plate. B and C show a photograph at close range of the pegs in the micro plate.

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After 3 min operation of the Terminator the homogenates are centrifuged in a micro plate centrifuge and the supernatant samples are used for IEF. 3. Pepper seeds are put in the wells of a 96-well micro plate, 200 μL DNA extraction buffer is added, and the seeds are disrupted and fully homogenized using the Terminator. After 3 min operation of the Terminator the homogenates are centrifuged in a micro plate centrifuge and 90 μL of each of the individual supernatants are brought (taking care not to disturb the pellets) in another micro plate. 90 μL PP buffer is added to the supernatants and the micro plate is mixed using an orbital shaker for 3 min at moderate speed. Then the plate is centrifuged at 4°C, 1,300g, for 10 min with moderate deceleration.

Of the resulting supernatants 90 μL is brought to a new micro plate and 90 μL ice-cold isopropanol (–20°C) is added to the supernatants. The micro plate is briefly shaken using the orbital shaker at medium speed and then centrifuged at 4°C, 1,300g for 10 min with moderate deceleration. The resulting supernatant is discarded and the pellet dried at 65°C for 20 min. To the pellet 100 μL TE-buffer is added, and the resulting DNA solution is stored at 4ºC. For long term storage –20°C is recommended.

3.2. Isoelectric Focusing (IEF) 1. For IEF of corn and sunflower seed extracts, gels of size 260 × 188 mm, and thickness of 0.2 mm are made fresh by combining the following solutions: 9.5 mL 16% glycerol, 2.0 mL acrylamide/bis acrylamide solution, 1.0 mL SinuLytes 3-7, 12 μL TEMED, 35 μL 10% ammonium persulfate. After swirling the solution the gel was poured. 2. The gel is divided in two fields using electrode paper wicks on the long sides and in the middle of the gel. In this way 96 samples can be run with a running distance of 5 cm. 3. Prefocusing is carried out at settings 600 Volts, 60 mA, 12 Watts for 75 Volthours 4. The corn and sunflower seed extracts (8 μL per sample) are applied to the horizontal IEF gel in the range 3–7 using the 96-well applicator strip. 5. Focusing is carried out in two runs. Run 1 with 200 Volts, 60 mA, 12 Watts for 50 Volthours and run 2 at settings 1000 Volts, 60 mA, 12 Watts for 1,000 Volthours. 6. After focusing, proteins are fixed in TCA solution for 10 min, stained with CBB solution for 10 min, and then the gel is destained several times for 10 min until the background is clear. 7. For ADH IEF of tomato seed extracts gels of size 260 × 188 mm, and thickness of 0.2 mm are made fresh by combining the following solutions: 9.5 mL 16% glycerol, 2.0 mL acrylamide/bis acrylamide , 1.0 mL SinuLytes 3-10, 12 μL TEMED, 35 μL l10% ammonium persulfate. After swirling the solution the gel was poured.

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8. The gel is divided in two fields using electrode paper wicks on the long sides and in the middle of the gel. In this way 96 samples can be run with a running distance of 5 cm. 9. Prefocusing is carried out at settings 600 Volts, 60 mA, 12 Watts for 75 Volthours 10. The tomato seed extracts (8 μl per sample) are applied to the horizontal IEF gel using the 96-well applicator strip. 11. Focusing is carried out in two runs. Run 1 with 200 Volts, 60 mA, 12 Watts for 50 Volthours and run 2 at settings 1000 Volts, 60 mA, 12 Watts for 700 Volthours. 12. The ADH staining solution is prepared just before the end of the focusing. Tris-HCl buffer is brought to 37°C. 5 mL ethanol, 0.05 g NAD, 0.05 g MTT and 0.01 g PMS are added, and dissolved by mixing. The gel is stained at 37°C until the bands can be clearly visualized (5–10 min). After removal of the stain, the gel is destained 10 min in 2% (v/v) acetic acid and then rinsed 10 min with water.

3.3. Inter-SSR PCR 1. The PCR mix is prepared just before PCR by adding the reaction mixture components together, including 3 μL of the DNA solution gained by extraction under Section 3.2. 2. The PCR mix is added to the PCR plates and PCR is carried out with the following program steps: an initial 5 min step at 94ºC, the 40 cycles of 0.3 min at 94°C, 0.45 min at 55°C, and 2 min at 72°C, which is followed by a final step of 5 min at 72°C.

3.4. DNA Electrophoresis 1. For horizontal electrophoresis of pepper ISSR PCR fragments, gels of size 260 × 188 mm and thickness of 0.2 mm are used. The following gel solution is used to pour the gels: 6.25 mL gel buffer, 9.5 mL 20% glycerol, 2.08 mL acrylamide/bis acrylamide 29: 1 solution, 0.02 mL TEMED, 0.14 mL 10% ammonium persulfate. The gel is poured and used the same day or stored at 4°C (for maximally 5 d). 2. To the PCR mix 4 μL of SB is added and of these samples, 4 μL is pipetted into the in-gel wells of the gel. 3. The gel is divided in two fields using electrode paper wicks on the long sides and in the middle of the gel. In this way 96 samples can be run with a running distance of 5 cm. 4. Electrophoresis is carried out at 15°C at power settings of 600 V, 40 mA and 24 W for 75 min 5. After electrophoresis the gel is incubated in fix solution for at least 3 min and then rinsed with water for 30 s. There after the gel is incubated in staining solution for 20 min, washed in water for 1 min, and then the DNA fragments are visualized by adding developer solution. The developer solution is refreshed after 1 min. The time for development is 3–5 min depending on the amount of DNA present

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4. Notes 1. Before operation of the Terminator, the head with the 96 pegs must be placed. It must be assured to place the head carefully, by lowering the Terminator head slowly above the Terminator base plate. Then it is slowly lowered to assure that the pegs fit into the wells of the micro plate. After operation, the head must be removed carefully to avoid sample going from one well to another. 2. The Voltage to be applied for operation of the Terminator must be determined empirically. The knob of the variable power supply must be turned slowly to increase the Voltage until the head starts clearly vibrating and macerating the tissue. This generates quite some noise by the pegs that hit the bottom of the plate. If the tissue is disrupted—this may take only a few seconds—the noise decreases, and homogenization can continue at a constant voltage somewhere between 90 and 130 V. The time needed for complete homogenization may also be determined empirically but is no more than 3 min. 3. Cleaning and maintenance of the Terminator is very simple. After operation, the Terminator head can be cleaned by spraying the pegs with water using a siphon and then the pegs can be dried on the air. The head can be placed with the pegs on a tissue. When kept clean, the Terminator needs no maintenance. Several Terminators operate now for more than 10 yr in several labs without maintenance. For thorough cleaning and decontamination the 96-peg plate can be detached from the vibrator head by turning the vibrator head counter clockwise. 4. Micro plates from Costar are used. However, as standard 96-well flat-bottom micro plates are produced worldwide using the same format, the Terminator is compatible with 96-well flat-bottom micro plates of all major suppliers. 5. SinuLytes are used as ampholytes in IEF gels. Alternatively, other sources of ampholytes are available. But SinuLytes are superior in our hands because of a low molecular weight (average molecular weight is between 400 and 700 Dalton) of the many amphoteric compounds, which means fast fixing, staining, and destaining of gels. In addition, high buffer capacity and solubility at pI, even conductivity along the gel and linear pH results in superior performance. 6. If one views the operation of the Terminator, one may easily suspect (at first sight) that cross-contamination (sample of one well contaminates sample of another well) is likely. However, extensive experiments to study possible cross-contamination were carried out. One may also view Fig. 3B. The single-banded pattern does not contain a trace of other bands that are present in other lanes. Further, for making the image of Fig. 4, pepper seeds were placed in such a way in the micro plates that each variety lacking the intense band (see Fig. 4) is surrounded by seeds of a variety that has the intense band. In this way cross-contamination would become easily visible from the resulting DNA banding pattern. This shows that

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Fig. 3. Images of isoelectric focusing gels showing analysis of homogenates prepared using the Terminator. A CBB stained banding pattern of corn seed extract focused in the pH range 3–7. The image shows genetic variability of the corn seed storage proteins. B CBB stained banding pattern of sunflower seed extract focused in the pH range 3–7. The image shows genetic variability of the sunflower seed storage proteins C ADH stained banding pattern of tomato seed extract in the pH range 3–10. The image shows the two known variants of the dimeric enzyme alcohol dehydrogenase from tomato seeds.

Fig. 4. Banding pattern resulting from ultra-thin layer electrophoresis of PCR samples using pepper seed DNA as template. One anchored primer is used for PCR. Pepper varieties were taken that differ in presence of an intense PCR band. Prior to homogenization using the Terminator, the pepper seeds were put in the micro plate in such a way that each seed having the band was surrounded by a seed lacking the band. The variable band is easily visible in the rectangle on the left in the image. An exploded view is made in the right at the bottom. The white square indicates the area of the gel that was eluted for further PCR (see Note 6).

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no cross-contamination occurred. To exclude even traces of cross-contamination, the gel area indicated with a white square in Fig. 4 was eluted and the sample was used for PCR. DNA electrophoresis showed no trace of DNA at that particular place in the gel. 7. Several types of cork borers are commercially available to cut round parts from large seeds such as corn, sunflower, and squash. Also leaf samples can best be taken using a cork borer or a puncher that cuts small leaf discs. It works best to cut discs of similar size as the bottom of the micro plate wells. First put the discs in the wells and then add the extraction fluid. In this way the leaf tissue can be optimally homogenized.

References 1. van den Berg, B.M. (1998) Isoelectric focusing in the vegetable seed industry. Electrophoresis 19, 1780–87. 2. van den Berg, B.M. and Tamboer, J.H.A. (1992) The terminator, an apparatus for simultaneous homogenization of 96 small seeds individually. Electrophoresis 13, 9, 10. 3. van den Berg, B.M. (1997) Horizontal ultrathin-layer multi-zonal electrophoresis of DNA: an efficient tool in analysis of PCR fragments. Electrophoresis, 18, 2861–64. 4. van den Berg, B.M., Burg, H.C.J., Tamboer, J.H.A., and Grapendaal, B. (1992) Equipment for rapid homogenization of high numbers of plant tissue for electrophoretic analysis of proteins. Electrophoresis 13, 76–81. 5. van den Berg, B.M. and Gabillard, D. (1994) Isoelectric focusing in immobilized pH gradient of melon (Cucumis melo L.) seed protein: methodical and genetic aspects and application in breeding. Electrophoresis 15, 1541–51. 6. van den Berg, B.M. (1990) Inbred testing of tomato (Lycopersicon esculentum) F1 varieties by ultrathin-layer isoelectric focusing of seed protein. Electrophoresis 11, 824–29. 7. van den Berg, B.M. (1991) A rapid and economical method for hybrid purity testing of tomato (Lycopersicon esculentum L.). F1 hybrids using ultrathin-layer isoelectric focusing of alcohol dehydrogenase variants from seeds. Electrophoresis 12, 64–9. 8. Zietkiewicz, E., Rafalski, A. and Labuda, D. (1994) Genome fingerprinting by simple sequence repeat (SSR)-anchored polymerase chain reaction amplification. Genomics 20, 176–83.

15 Isolation of Mitochondria from Plant Cell Culture Etienne H. Meyer and A. Harvey Millar

Summary Mitochondria carry out a variety of important processes in plants. Their major role is the synthesis of ATP through the coupling of a membrane potential to the transfer of electrons from NADH to O2 via the electron transport chain. The NADH is generated by the oxidation of organic acids via the tricarboxylic acid cycle. However, mitochondria also perform many important secondary functions such as synthesis of nucleotides, amino acids, lipids, and vitamins. Mitochondria contain their own genome and undertake transcription and translation by some unique mechanisms; they actively import proteins and metabolites from the cytosol, are involved in programmed cell death processes in plants, and respond to cellular stress conditions. To understand the extent and mechanisms of mitochondrial functions in plants and the way in which their functions are perceived by the nucleus requires detailed information about the protein components of these organelles. Isolation of mitochondria to identify their proteomes and the changes in these proteomes during development and environmental stresses is growing area of research. In this chapter we provide a useful method for the isolation of mitochondria from plant cell culture using a gentle method of cell disruption based on protoplasts isolation that provides relatively high mitochondrial yields.

Key Words: Cell fractionation; mitochondria; percoll density gradients; protoplasts isolation.

1. Introduction Plant mitochondria play an important role in plant metabolism. They provide energy in the form of ATP, but also they provide a large number of metabolic precursors in the form of tricarboxylic acids to the rest of the cell for nitrogen assimilation and biosynthesis of amino acids. They play key roles in plant From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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development, fertility, and susceptibility to disease. Between 1,000 and 1,500 proteins are expected to be found in this cellular compartment and currently many hundreds of these proteins have been identified by proteomics. Studies of the plant mitochondria proteome require isolation methods that avoid rupture of mitochondrial membranes. These methods should be able to eliminate most of the cellular contaminant released after disruption of the plant cell. Several extensive methodology reviews (1–3) and more specific methodology papers (4–6) are already available on plant mitochondrial purification. All these methods are based on a cell disruption by grinding. Here we describe a gentle method based on protoplasts isolation. Protoplasts are plant cells after removal of the cell wall. This cell wall is made of fibrils of cellulose embedded in a matrix of several other kinds of polymers such as pectin and lignin. This rigid structure can be digested using two enzymatic activities, cellulase will digest the cellulose and pectolyase will break the intercellular pectin bonds. The resulting protoplasts are highly fragile and can easily be broken by filtration or homogenization. Protoplasts can be isolated from virtually any plant tissue. Plant organs give low yields of protoplasts, whereas cell cultures are an excellent starting material for protoplast isolation. The method we describe here was optimized for the purification of mitochondria from plant cell culture.

2. Materials 1. Plant cell material for protoplasts: Depending on the growth rate of the plant culture, 5–7 day old cell culture should be used. This culture should be sterile to avoid fungal and bacterial contamination. 2. Enzyme buffer: 0.4M mannitol, 0.7 g/L MES-KOH pH 5.7. Just before use 0.4% (w/v) of cellulase and 0.05% (w/v) of pectolyase are added (see Note 1). This buffer has to be prepared just before use. 3. Disruption buffer: 0.4M sucrose, 3 mM EDTA, 50 mM Tris-HCl, pH 7.5, 0.1% BSA, 2 mM DTT (Dithiothreitol) which is added just prior the disruption. The osmoticum (sucrose) maintains the mitochondria structure and prevents physical swelling and rupture of membranes, the buffer (Tris) prevents acidification from the contents of ruptured vacuoles, the EDTA inhibits the function of phospholipases and various proteases, the BSA will remove free fatty acids and the reductant (DTT) prevents damage from oxidants present in the tissue or produced on homogenization. This media can be freshly prepared, stored overnight at 4°C or frozen at –20°C and stored for many weeks (see Note 2). 4. Wash buffer: 0.3 M sucrose, 1 mM EDTA, 10 mM MOPS-KOH, pH 7.2. This buffer is then used for resuspension of organelle pellets, as the base media for Percoll gradients and for washing purified organelle pellets. This media can be freshly prepared, stored overnight at 4°C or frozen at –20°C and stored for many weeks (see Note 2).

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5. Percoll gradient solutions: Gradients of Percoll (Pharmacia, Uppsala, Sweden) (see Note 3) are prepared in the wash buffer on the day of use. This is aided by making a 2× wash buffer and adding Percoll and distilled water to make 1× wash buffer with the appropriate percentage of Percoll required. The discontinuous step gradient was cast using a simple inverted syringe bodies (see Note 4).

3. Methods 3.1. Protoplasts Isolation The cell culture (5–7 day-old cells) is filtered through two layers of muslin. The cells are resuspended in the enzyme buffer (a maximum of 500 g of cells per L of buffer) and incubated for 3 h in the dark under low agitation (45 rpm) at 25°C. Then the protoplasts are washed twice to remove all traces of the digestion enzymes. The suspension is centrifuged (for 10 min at 800g) and the pelleted protoplasts are resuspended in enzyme buffer without enzyme (see Note 5). After the second wash, the protoplast pellet is resuspended in cold disruption buffer. The following steps should be done either in a cold room or on ice using 4ºC cooled glassware and centrifuge tubes (see Note 2). 3.2. Protoplasts Disruption The protoplasts can be disrupted by filtration through Nylon meshes. The suspension is filtrated successively through three different Nylon meshes (100 μm, 75 μm, and 30 μm holes). Alternatively, the protoplasts can be broken by homogenization in a Dounce homogeniser (or potter) (see Note 6). We recommend checking the digestion as well as the disruption by optical microscopy (see Note 7). 3.3. Differential Centrifugation to Obtain a Crude Organelle Pellet 1. Transfer filtered homogenate into 50, 250, or 500 mL centrifuge tubes, depending on the volume of the preparation, and centrifuge in a precooled rotor for 5 min at ∼3,000g in a fixed angle rotor in a preparative centrifuge at 4°C. 2. Decant supernatant gently into another set of centrifuge tubes taking care not to transfer the pellet material which contains plastids, nuclei and cell debris. Centrifuge supernatant for 15 min at ∼18,000g and the resulting high speed supernatant is discarded. The tan, yellow, or green coloured pellet in each tube contains an unwashed crude organelle pellet. 3. Resuspend the pellet in 2–10 mL of wash medium with the aid of a clean, soft bristle paint brush.

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Fig. 1. Percoll gradient purification of plant mitochondria. (A) Three-step Percoll gradient for the purification of Arabidopsis mitochondria made from 5 mL of 50% Percoll, 25 mL of 25% Percoll, and 5 mL of 18% Percoll solution from bottom to top. This gradient was centrifuged at 40,000g in a fixed angle rotor for 45 min. Amyloplast envelopes are concentrated in the 18 to 25% interphase (“a”), the fraction containing mitochondria in the 25–40% interphase (“m”). (B) One-step Percoll gradient made with 35 mL of 28% Percoll. The fraction containing mitochondria from the three-step Percoll gradient was loaded on this gradient which was then centrifuged at 40,000g in a fixed angle rotor for 45 min. Mitochondria (“m”) are present on top of the gradient whereas peroxisomes and other contaminants (“p”) are located in the bottom part of the gradient.

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3.4. Density Gradient Puriﬁcation of Mitochondria The crude mitochondrial preparation described above is contaminated by thylakoid or amyloplast membranes, peroxisomes and endoplasmic reticulum. Further purification is carried out using Percoll (Pharmacia, Uppsala, Sweden) density gradients. 1. Layer washed mitochondria, from up to 80 g of etiolated plant tissue or up to 40 g green plant tissue, over 35 mL of a 18–25–40% step gradient (from bottom to top : 5 mL of 50%, 25 mL of 25%, 5 mL of 18% Percoll in wash buffer) in a 50 mL centrifuge tube (see Note 2). 2. Centrifuge at ∼40,000g for 45 min in a fixed angle rotor of a preparative centrifuge without braking on the deceleration. 3. After centrifugation, mitochondria form a white-brown band in the bottom part of the gradient (Fig. 1). Aspirate the mitochondria with a Pasteur pipet avoiding collection of the yellow or green plastid fractions. Dilute suspension with at least four volumes of standard wash medium and centrifuge at ∼18,000g for 15 min in 50 mL tubes. 4. The resultant loose pellets is resuspended in a small amount of wash medium and loaded on top of a 28% Percoll continuous gradient (35 mL of 28% Percoll in wash buffer in a 50-mL centrifuge tube). 5. Centrifuged at ∼40,000g for 45 min in an angle rotor of a preparative centrifuge without braking on the deceleration. 6. After centrifugation the mitochondria form a white band in the top part of the gradient whereas contaminants such as peroxisomes are located in the bottom part of the gradient (Fig. 1). Aspirate the mitochondria with a Pasteur pipet and dilute them with at least four volumes of wash buffer and centrifuge again at ∼18,000g for 15 min in 50-mL tubes. 7. Remove the supernatant and resuspend the pellet in wash buffer. Centrifuge again at ∼18,000g for 15 min. Resuspend the mitochondrial pellet in wash medium at a concentration of 5–20 mg mitochondrial protein/mL. This can be determined using a Bradford or Lowry assay. 8. In our hands 100 g of cell culture yields 95 g of protoplasts and 15 mg of purified mitochondria. 9. Once isolated by density gradient purification, plant mitochondria can be kept on ice for 5–6 h without significant losses in membrane integrity and respiratory function. Longer-term storage of mitochondria can be achieved by rapid-freezing of mitochondrial samples in liquid N2 . Frozen samples can be then kept at –80°C.

4. Notes 1. At these concentrations of enzymes, more than 95% of the cells will be converted in protoplasts after 3 h. Increasing the concentrations may result in a faster

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digestion but also a lot of protoplasts could break due to a longer incubation in the buffer. 2. Preparation of mitochondria should be undertaken as quickly as is possible and without samples warming above 4°C or storage for extended periods between centrifugation runs. So we highly recommend using chilled buffers. The time between homogenization and preparation of the washed crude pellet is the most critical for ensuring integrity and high yield. 3. The colloidal silica sol, Percoll, allows the formation of iso-osmotic gradients and through isopycnic centrifugation facilitates a range of methods for the density purification of mitochondria. The most common method is the sigmoidal, selfgenerating gradient obtained by centrifugation of a Percoll solution in a fixedangle rotor. The density gradient is formed during centrifugation at >10,000g due to the sedimentation of the poly-dispersed colloid (average particle size 29 nm diameter, average density = 2.2 g/mL). The concentration of Percoll in the starting solution and the time of centrifugation can be varied to optimise a particular separation 4. Step gradients of Percoll are often used as these aids the concentration of mitochondria fractions on a gradient at an interface between Percoll concentrations. Step gradients can easily be formed by setting up a series of inverted 20-mL syringes (fitted with 19-gauge needles) strapped to a flat block of wood, clamped to a retort stand over a rack at an angle of 45° containing the centrifuge tubes (Fig. 2). The needles are lowered to touch the bevels against the inside, lower edge of the tubes. The step gradient solutions are then added (from bottom to top) to the empty inverted syringe bodies and each allowed to drain through in turn before the addition of the next step solution.

Fig. 2. Making density gradient. Home-made apparatus for discontinuous gradients, see explanation in Note 4

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5. The protoplasts are very fragile. Thus the agitation should be very gentle (45 rpm) on an orbital shaker. Also, the resuspension of the pelleted protoplasts should be as soft as possible to avoid disruption of protoplasts. We recommend resuspending the pellet by slowly swirling the tube. 6. These methods of disruption are dependant on the size of the cells. Small cells (diameter below 10 μm) will not be disrupted. Mesh with smaller holes should then be used. The disruption of large cells will release a lot of membrane fragment which will block the holes of the 10-μm mesh. Then the filtration through the 10-μm mesh will be replaced by a second filtration through the 30-μm mesh. 7. The digestion has to be checked by optical microscopy after three hours. A drop (approx 50 μL) is largely sufficient. If a lot of nondigested cells remain (nonround cells), incubate the suspension a longer time in the enzyme medium. The disruption should also be checked to ensure that all the protoplasts are broken. Unbroken protoplasts will be pelleted during the low-speed centrifugation and lost. If some unbroken protoplasts remain, repeat the filtration or homogenisation step.

References 1. Neuburger, M. (1985) Higher-Plant Cell Respiration, vol. 18 (Douce, R., Day, DA, ed.), pp. 7–24, Springer-Verlag, Berlin. 2. Douce, R. (1985) Mitochondria in higher plants: Structure, function and biogenesis, American Society of Plant Physiologists, Academic Press, Orlando, Florida. 3. Millar, A. H., Liddell, A., and Leaver, C. J. (2001) Isolation and subfractionation of mitochondria from plants. Meth. Cell Biol., 65, 53–74. 4. Neuburger, M., Journet, E. P., Bligny, R., Carde, J. P., and Douce, R. (1982) Purification of plant-mitochondria by isopycnic centrifugation in density gradients of percoll. Arch. Biochem. Biophys. 217, 312–23. 5. Leaver, C. J., Hack, E., and Forde, B. G. (1983) Protein-synthesis by isolated plant-mitochondria. Meth. Enzymol. 97, 476–84. 6. Day, D. A., Neuburger, M., and Douce, R. (1985) Biochemical-characterization of chlorophyll-free mitochondria from pea leaves. Aust. J. Plant Physiol. 12, 219–228.

16 Isolation and Preparation of Chloroplasts from Arabidopsis thaliana Plants Sybille E. Kubis, Kathryn S. Lilley, and Paul Jarvis

Summary A major area of research in the postgenomic era has been the proteomic analysis of various subcellular and suborganellar compartments. The success of these studies is to a large extent dependent upon efficient protocols for the preparation of highly pure organelles or suborganellar components. Here we describe a simple, rapid, and low-cost method for isolating a high yield of Arabidopsis chloroplasts. The method can readily be applied to wild-type plants and different mutants, and at different developmental stages ranging from 10-day-old seedlings to rosette leaves from older plants. The isolated chloroplast fraction is highly pure, with immunologically undetectable contamination from other cellular organelles. Chloroplasts isolated using the method described here have been successfully used for proteomic analysis, as well as in studies on chloroplast protein import and other aspects of chloroplast biology.

Key Words: Arabidopsis thaliana; chloroplast isolation; chloroplast proteomics; organelle isolation; Percoll gradient; plastids; polytron homogenizer.

1. Introduction Chloroplasts belong to a diverse group of organelles called plastids (1,2). Plastids are ubiquitous in plants and algae, and perform numerous essential functions including important steps in the biosynthesis of amino acids, lipids, nucleotides, hormones, vitamins, and secondary metabolites, as well as oxygenic photosynthesis (2,3). In the latter process, energy from sunlight is converted into usable chemical bond energy, and the associated redox reactions lead to the generation of oxygen from water. Chloroplasts are therefore From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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important sites for the production of organic matter and oxygen, and so provide the fuels essential for all higher forms of life (4). Completion of the genome sequencing projects for Arabidopsis, rice and other species, and the development of efficient methods for routine protein identification by mass spectrometry, have enabled numerous largescale proteomic studies. Because of the dynamic range limitations associated with analyses on highly complex mixtures (i.e., the tendency of abundant proteins to mask the presence of less abundant proteins), these studies have tended to focus on isolated subcellular components. In plants, chloroplasts have received considerable attention in this regard (5–8). The uses of such proteomic studies are several-fold: they can confirm the expression and structure of genes predicted based on genome sequence analysis in silico; they can provide information on subcellular and suborganellar protein localization; and they can even be used to estimate the relative abundance of different proteins. Information of this nature is particularly important, because it has been estimated that up to 50% of the proteins encoded by the >26,000 genes in the Arabidopsis genome are of unknown function (9). Proteomics is one of the tools being used to address this deficiency. Most proteins targeted to chloroplasts possess a cleavable, amino-terminal targeting signal called a transit peptide (10,11). Using computer programs (e.g., TargetP) to detect the presence of a transit peptide, it has been estimated that ∼4,000 proteins are targeted to chloroplasts in Arabidopsis (9). Unfortunately, these in silico methods are not 100% reliable (12), and so the only truly dependable method for the determination of protein localization is laboratory experimentation. However, there are presently less than 700 entries in a database of experimentally determined chloroplast proteins (13), clearly indicating a need for further studies. Furthermore, several recent reports have indicated that protein targeting to chloroplasts is not as simple as was once thought. In a large-scale study of the Arabidopsis chloroplast proteome, only ∼60% of the proteins identified were predicted to have a transit peptide (6,14). Of the remainder, many appeared to have a signal peptide for ER translocation, or no cleavable targeting sequence at all. Intriguingly, direct evidence for a protein transport pathway to chloroplasts through the ER and Golgi has now been presented (15), and the targeting of proteins lacking a cleavable peptide has been described in some detail (16,17). These data demonstrate that transit peptide prediction in silico cannot provide a complete picture of the chloroplast proteome. The existence of dual-targeted proteins (e.g., proteins targeted to mitochondria or the ER as well as chloroplasts) adds an additional level of complexity (18,19), further emphasizing the need for experimental determination of protein localization.

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A number of chloroplast proteomic studies have already been described. In addition to the whole chloroplast study mentioned earlier (14), several reports have described the proteomes of individual suborganellar compartments of the chloroplast: e.g., the thylakoid lumen (20,21), the envelope membrane (22,23), the stroma (24), and the lipid-containing structures called plastoglobuli (25). In addition, an analysis of plastids in dark-grown plants, called etioplasts, revealed a proteome consistent with what one would expect of plastids in heterotrophic tissue, along with some novel functions (26). As well as studies that simply catalogue the proteins present in a particular subcellular or suborganellar compartment, comparative proteomics has been employed with considerable success. For example, the chloroplasts from mesophyll cells and bundlesheath cells of maize, a C4 plant, were recently compared (27). The data not only revealed differential accumulation of carbon metabolism enzymes consistent with the C4 photosynthetic mechanism, but also shed light on how other plastidic functions are distributed between the two cell types. In another example, chloroplasts isolated from Arabidopsis mutants lacking different protein import receptor isoforms were compared with wild-type chloroplasts (28,29). Different groups of chloroplast proteins were found to be selectively deficient in different receptor mutants, indicating that the different receptor isoforms likely possess a degree of preprotein recognition specificity (10,11). These various studies have demonstrated the utility and value of chloroplast proteome analysis, and it is anticipated that proteomics will continue to form an essential component of chloroplast research in the future. The chloroplast isolation procedure described in technical detail here, and previously (30), has been successfully used in proteomic studies (28,29), as well as in other research on chloroplast biology (31–33).

2. Materials 2.1. Growth of Arabidopsis Seedlings 1. Seeds, stored in a 1.5-mL microfuge tubes (with a small hole in the lid to allow evaporation of any residual moisture) at room temperature. 2. 70% (v/v) ethanol containing 0.05% (v/v) Triton X-100 (Sigma-Aldrich Ltd., Poole, UK), in a Duran bottle at room temperature. For 200 mL, mix 140 mL 100% ethanol, 60 mL sterile deionized water, and 100 μL Triton X-100. 3. 100% ethanol, in Duran bottle at room temperature. 4. Laminar flow hood (e.g., Model P5HB, Bassaire Ltd., Southampton, UK). 5. Circular filter papers, 9 cm in diameter (Whatman, Banbury, UK; or Fisher Scientific, Loughborough, UK). 6. Industrial methylated spirit (IMS). 7. Orbital shaker (e.g., Model S01, Stuart Scientific, Stone, UK).

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8. Petri dishes, 9 cm in diameter (e.g., Bibby Sterilin Ltd., Stone, UK). 9. Murashige and Skoog (MS) medium: MS salt and vitamin mixture (Sigma), 0.5% (w/v) sucrose, and 0.6% (w/v) agar. Sterilize the medium in an autoclave (15 min, 121°C, 15 psi), cool to 50°C in a water bath, and then pour into Petri plates to a depth of ∼3–4 mm in a laminar flow hood (400 mL medium is sufficient for ∼15–20 plates). Allow the plates to dry for 1 h in the hood before replacing the lids. Prepoured plates can be stored at 4°C, up-side down, sealed in a plastic bag for up to 1 month. 10. Leukopor tape (Beiersdorf AG, Hamburg, Germany) or equivalent (e.g., Micropore tape, 3M, Bracknell, UK). 11. Refrigerator or cold-room (4°C). 12. Plant tissue culture chamber (e.g., Model CU-36L5, Percival Scientific Inc., Perry, Iowa) set at 20°C, providing 100–120 μmol/m2 /s white light with a longday cycle (16-h-light/8-h-dark).

2.2. Chloroplast Isolation These materials are sufficient for one isolation on a single plant sample. If multiple samples (e.g., different genotypes) are to be analyzed, additional materials will be required (i.e., in points 1, 3, 6, 7, and 9 below). 1. For one isolation procedure, 25–40 Petri plates of 10-day-old plants, each plate containing ∼150–200 seedlings as shown in Fig. 1B (see Notes 1, 2). 2. Two ice buckets, containing ice. 3. Two 1-L beakers, one 50-mL beaker, measuring cylinders, and one funnel. 4. Polytron; e.g., Kinematica Model PT10-35 (Kinematica AG, Littau, Switzerland), with a large rotor (PTA 20 S) and speed set to 4 on scale of 11 (see Note 3). 5. Cold-room (4°C). 6. Miracloth (Calbiochem Ltd., Nottingham, UK); two squares of about 15 × 15 cm. 7. Two 30-mL Nalgene tubes and one 250-mL Nalgene tube (Fisher). 8. Percoll (Amersham Biosciences, Little Chalfont, UK); an opened bottle can be stored at 4°C for several months. 9. Continuous Percoll gradient. Before use, make up 26 mL of gradient mixture as follows: 13 mL Percoll, 13 mL 2× chloroplast isolation buffer (see below), and 5 mg glutathione (roughly, the tip of a small spatula). Mix the components together in a 30-mL Nalgene tube, ensuring that the glutathione is completely dissolved. Precentrifuge in a fixed angle rotor at 43,000gmax for 30 min (brake off) at 4°C; this is equivalent to 19,000 rpm in an SS-34 rotor in a Sorvall RC6 centrifuge (Kendro, Asheville, North Carolina), with acceleration set to 7 and deceleration set to 2. Gradients can be prepared the day before, and then stored overnight at 4°C. 10. Chloroplast isolation buffer (CIB): 0.3M sorbitol, 5 mM MgCl2 , 5 mM EGTA, 5 mM EDTA, 20 mM HEPES/KOH pH 8.0, 10 mM NaHCO3 . This is prepared as a 2× CIB stock (see Notes 4, 5). The final pH of the solution should be 8.0.

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Fig. 1. Different steps of the chloroplast isolation procedure. (A) An MS agar plate carrying Arabidopsis seeds sown at an appropriate density (∼150 seeds per plate) for use in chloroplast isolation after growth for 10–14 d. (B) A plate similar to that shown in A, after the plants have been allowed to grow for 14 d in a tissue culture cabinet. (C, D) The Arabidopsis seedlings are harvested from the plates by hand, taking care not carry over any of the agar medium. (E) The Arabidopsis tissue is transferred to a 50-mL beaker containing cold chloroplast isolation medium, and then disrupted using five consecutive rounds of homogenization using a polytron blender. (F) Following filtration through Miracloth, the homogenate is loaded onto a preformed continuous Percoll gradient. (G) After centrifugation of the loaded Percoll gradient, two green bands are apparent: the upper band contains broken material, whereas the lower band contains intact chloroplasts.

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11. HEPES-MgSO4 -sorbitol (HMS) buffer: 50 mM HEPES/NaOH pH 8.0, 3 mM MgSO4 , 0.3M sorbitol (see Note 6). The final pH of the solution should be 8.0.

2.3. Establishing Yield and Intactness of Chloroplasts 1. Hemocytometer with a 0.1 mm deep counting chamber and a ruling pattern of 1/400 mm2 (e.g., Improved Neubauer BS748, Hawksley Technology, Lancing, UK). 2. Cover glass (e.g., 22 × 22 mm, No.1, Chance Propper Ltd., Warley, UK). 3. Phase-contrast microscope (e.g., Carl Zeiss AG, Oberkochen, Germany). 4. HMS buffer (see Section 2.2.). 5. Tissue paper (e.g., Kimcare, Kimberly-Clarke Europe Ltd., Reigate, UK).

2.4. Preparation for Proteomics 1. Ice bucket containing ice, with lid. 2. Lysis buffer (see Table 1). Recommended detergent components: ASB14 (amidosulfobetaine-14) (Calbiochem Ltd.); CHAPS (3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate) (Sigma-Aldrich Ltd.); SB3-10 Table 1 Lysis buffers Buffer 1. Standard buffer

2. Thiourea buffer

3. ASB-14 buffera

4. SDS bufferb

a

Component CHAPS Urea Tris-HCl, pH 168–169 Magnesium acetate CHAPS Urea Thiourea Tris-HCl, pH 8.0–9.0 Magnesium acetate ASB-14 Urea Thiourea Tris-HCl, pH 8.0–9.0 Magnesium acetate SDS Tris-HCl, pH 8.0–9.0 Magnesium acetate

Concentration 4% (w/v) 8M 10–30 mM 5 mM 4% (w/v) 7M 2M 10–30 mM 5 mM 2% (w/v) 7M 2M 10-30 mM 5 mM 2% (w/v) 10-30 mM 5 mM

ASB-14 can be substituted with NP40, SB3-10 or various other sulfobetaine-derived detergents. b SDS-containing solutions must be diluted to a final concentration of 0.2% or less before successful isoelectric focusing can take place.

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(3-(decyldimethylammonio)propanesulfonate inner salt) (Sigma-Aldrich Ltd.); Nonidet P40 (NP40) substitute ([octylphenoxy]polyethoxyethanol) (USB Corp., Cleveland, Ohio). Hand-held, plastic pestles for use with 1.5-mL microfuge tubes (e.g., pellet pestle, blue polypropylene, Sigma-Aldrich Ltd.). Bench-top microcentrifuge (e.g., Eppendorf 5415D, Eppendorf UK Ltd., Cambridge, UK). Protein concentration estimation kit (e.g., DC Protein Assay, Bio-Rad Laboratories Ltd., Hemel Hempstead, UK; or PlusOne 2-D Quant, Amersham Biosciences). Plastic cuvets, 1 mL (e.g., Sarstedt Ltd., Leicester, UK). Spectrophotometer (e.g., Spectronic; Thermo Electron Corp., Waltham, Massachusetts).

3. Methods 3.1. Growth of Arabidopsis Seedlings 1. Transfer the appropriate amount of seeds (e.g., for 40 Petri plates each carrying ∼150–200 seeds, an amount equivalent to ∼240–320 μL will be required) into a sterile 1.5-mL microfuge tube, and add 1 mL of 70% (v/v) ethanol, 0.05% (v/v) Triton X-100 (see Note 7). 2. Shake the tube by hand to ensure that all seeds are suspended in the solution, and then place the tube, oriented horizontally, onto an orbital shaker. Shake at 250 rpm for 5 min. 3. Allow the seeds to settle at bottom of tube, remove the supernatant with a pipet, and then add 1 mL of 100% ethanol. Shake the tube by hand first of all, and then on the orbital shaker at 250 rpm for 10 min. 4. Meanwhile, switch on the laminar flow hood and sterilize the interior surfaces with IMS. Take an appropriate number of round filter papers (at least one per seed sample), and fold them in half to create a crease (this will facilitate seed sowing later). In the hood, soak (and sterilize) the filter paper(s) with IMS. Allow the filter paper(s) to dry. 5. Using a cut 1-mL Gilson pipet tip (lacking ∼5 mm from the fine end, to increase the aperture size), pipet the seeds onto the sterilized filter paper(s) and leave to dry. This takes about 15 min. 6. Sow seeds onto Petri plates containing MS medium. For 10- to 14-day-old plants, an appropriate density is ∼150–200 seeds/plate, as shown in Fig. 1A. (see Notes 7, 8). 7. Seal each plate with Leukopor tape. 8. Incubate plates upside down (to prevent condensation accumulating on the surface of the agar) at 4°C for at least 2 d (up to 4 d is possible) to break seed dormancy and synchronize germination. 9. Grow plants for 10–14 d in a plant tissue culture chamber. Plants grown for 14 d are shown in Fig. 1B.

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3.2. Isolation of Chloroplasts The isolation procedure should be started as early as possible in the morning. During the isolation procedure, plant material should kept at 4°C. The first steps (stages 2–4 below) can be carried out on the bench in the laboratory, but the isolation itself should be carried out at 4°C in the cold-room. The method below describes an isolation on a single plant sample. If multiple samples (e.g., different genotypes) are to be used, additional materials will be required (see Section 2.2). 1. Preparation, on the day before the isolation: Place a 200-mL aliquot of 2× CIB, a 50-mL aliquot of 2× CIB, a 50-mL aliquot of HMS buffer, and 200 mL deionized H2 O into the refrigerator or cold-room; in the morning, the frozen solutions should have thawed. Place all rotors in the cold-room to precool overnight. 2. Preparation, on the day of the isolation: Prepare CIB by adding 200 mL chilled sterile deionized H2 O to 200 mL thawed 2× CIB; mix well and keep on ice. Put 100 mL of the CIB into a 1-L plastic beaker and keep on ice. Place a second 1-L plastic beaker on ice, with funnel containing two layers of Miracloth. The 250-mL and 30-mL Nalgene tubes should also be placed on ice to precool. 3. Prepare a continuous Percoll gradient as described in Section 2.2 (point 9), and keep the tube on ice after precentrifugation (see Note 9). 4. Take plates out of plant tissue culture chamber and remove the Leukopor tape. Remove the seedlings from the medium by gently scraping them off with a gloved hand (see Fig. 1C, D), avoiding carry over of medium because this interferes with the isolation, and place them into the 100 mL CIB in the 1-L beaker on ice. 5. During homogenization, a total of 100 mL CIB is used per sample; this is used in five, consecutive rounds of homogenization, each one using 20 mL CIB (see Note 10). 6. Place 20 mL fresh CIB into the 50-mL plastic beaker, and then transfer the seedlings into the beaker. 7. Place the plant material under the rotor of polytron, and homogenize for 1-2 s (see Fig. 1E). The optimal conditions for the homogenization have to be established empirically (see Note 11). 8. Filter the homogenate through two layers of Miracloth into the 1-L beaker on ice. Gently squeeze the Miracloth around the plant material. 9. Place a second 20-mL aliquot of fresh CIB into 50-mL beaker, and return the plant material to the beaker. 10. Repeat points 7–9 until all five 20-mL aliquots of CIB have been used, and five rounds of homogenization and filtration have been completed. The plant material will gradually become disrupted during the procedure. 11. Transfer the pooled, filtered homogenate into the 250-mL Nalgene tube on ice, and centrifuge at 1,000gmax for 5 min (brake on) at 4°C; this is equivalent to 3,000 rpm in an SLA-1500 rotor in a Sorvall RC6 centrifuge, with both acceleration and deceleration set to 7).

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12. Pour off most of the supernatant, and resuspend the pellet in the residual ∼500 μL supernatant by rotating the tube on ice; do not resuspend by pipeting. 13. Transfer the resuspended homogenate onto the top of the preformed Percoll gradient, using a cut 1-mL Gilson pipet tip (lacking ∼5 mm from the fine end, to increase the aperture size). Pipet very slowly so as not to disturb the gradient (see Fig. 1F). 14. To separate the intact chloroplasts from broken chloroplasts and other debris, centrifuge in a swing-out rotor at 7,800gmax for 10 min (brake off) at 4°C; this is equivalent to 7,000 rpm in an HB-6 rotor in a Sorvall RC6 centrifuge, with acceleration set to 7 and deceleration set to 2. 15. After centrifugation, remove the tube carefully and place it on ice. The lower green band in the gradient contains intact chloroplasts, whereas the upper band contains broken chloroplasts (see Fig. 1G). Broken chloroplasts are removed and discarded by pipeting, and then the intact chloroplasts are recovered using a 1-mL Gilson pipet tip (cut at the end), and transferred into a precooled 30 mL Nalgene tube. The volume of recovered intact chloroplasts can range from 2 mL to 6 mL. 16. Add 25 mL HMS buffer to the chloroplasts and invert the tube carefully 2–3 times to wash off the Percoll. 17. Centrifuge the chloroplasts in a swing-out rotor at 1,000gmax for 5 min (brake on) at 4°C; this is equivalent to 2,000 rpm in an HB-6 rotor in a Sorvall RC6 centrifuge, with both acceleration and deceleration set to 7. 18. Gently pour off the supernatant, and then resuspend the chloroplasts in ∼150–400 μL fresh HMS buffer by rotating the tube on ice; do not resuspend by pipeting.

3.3. Establishing the Yield, Intactness and Purity of the Isolated Chloroplasts If necessary, the yield of chloroplasts and their intactness can be assessed as follows. Alternatively, for some applications it may be appropriate to proceed directly to downstream procedures (e.g., Section 3.4). 1. Add 5 μL of isolated chloroplasts to 495 μL of HMS buffer in a 1.5-mL microfuge tube, and then mix gently by flicking the tube to obtain a 1:100 dilution. 2. Pipet ∼60–80 μL of the diluted suspension onto the counting chamber of the hemocytometer, and place a cover glass on top. 3. Drain the excess suspension with tissue paper. 4. Count the number of chloroplasts in 10 different 1/400 mm2 squares (e.g., those on each diagonal line), using a phase contrast microscope with a 16× objective. The number of chloroplasts per square should average between 10 and 20. If too few or too many chloroplasts are present, adjust the dilution factor (point 1 above) accordingly and repeat the procedure. Intact chloroplasts appear round and bright green (see Fig. 2A), and under phase-contrast are surrounded by a bright halo of light.

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5. The concentration (number of chloroplasts per mL) is calculated as follows: n (the average number of chloroplasts per 1/400 mm2 square) × 25 (the total number of squares in the grid) × 100 (the dilution factor employed) × 104 (scaling factor needed to express the data per mL, because the volume above the 25 squares is only 0.1 μL). 6. Calculate the actual yield of chloroplasts by multiplying the concentration (number of chloroplasts per mL) by the volume of chloroplast suspension used in Section 3.2, point 18.

Samples prepared using the methodology described here are mostly intact, and exhibit minimal contamination from other cellular compartments. To illustrate these points, we analyzed typical chloroplast preparations by phasecontrast light microscopy (see Fig. 2A,B), by transmission electron microscopy (see Fig. 2C), and by immunoblotting using high-titre antibodies against components of various cellular organelles (see Fig. 3). Microscopic analysis did not reveal any evidence of significant contamination from other cellular organelles.

Fig. 2. Light and electron micrographs of isolated chloroplasts. (A, B) Chloroplasts isolated from 14-day-old Arabidopsis seedlings were analysed by phase-contrast light microscopy, at both low (A) and high (B) magnification, and the majority were adjudged to be intact. Size bars indicate 100 μm. (C) The integrity of the chloroplasts was confirmed by transmission electron microscopy, as described previously (30). No evidence of significant contamination of the chloroplast preparation with other organelles was observed using either method. Size bar indicates 2 μm.

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Immunologically, the chloroplasts contained undetectable contamination from all organelles tested: <1% from the plasma membrane, mitochondria and peroxisomes, and <2.5% from the nucleus (the slightly higher value here simply reflects the lower sensitivity of the antibody used) (see Fig. 3).

Fig. 3. Immunological estimation of contamination of the isolated chloroplasts. To assess for contamination of chloroplast preparations from other cellular compartments, immunoblot analyses were performed on dilution series of total protein extract (left) and isolated chloroplasts (right); both samples were derived from 10-day-old, wildtype Arabidopsis plants. The indicated amounts of each protein were loaded, and the blots were probed with antibodies against proteins from various different subcellular compartments, as indicated. Because approximately half of the protein in the total protein extract is chloroplast-derived, 2 μg total protein is approximately equivalent to 1% of 100 μg chloroplast protein, and 5 μg total protein is approximately equivalent to 2.5% of 100 μg chloroplast protein. Thus, the detection of a protein band using 2 μg (PMA2, PrxII F and catalase) or 5 μg (histone H3) of total protein, together with the failure to detect a band using 100 μg chloroplast protein, indicates that we have <1% or <2.5% contamination, respectively, of the chloroplast sample with proteins from other compartments. In fact, contamination was immunologically undetectable. The antibodies used have been described previously (29,34–36), or were obtained commercially (histone H3; Abcam, Cambridge, UK). Abbreviations used are as follows: PMA2 (plasma membrane H+ -ATPase 2), PrxII F (type II peroxiredoxin F), PSI-D (photosystem I subunit D), and SSU (small subunit of Rubisco).

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3.4. Preparation of the Chloroplasts for Proteomics 1. Chloroplast lysis and solubilization: Resuspend the intact chloroplast sample in a small volume of the appropriate lysis buffer (see Table 1). To ensure optimal lysis, a plastic hand-held pestle should be employed in conjunction with a 1.5-mL microfuge tube. Solubilization should be allowed to proceed on ice for 30 min, with frequent grinding of the chloroplast material using the pestle (see Note 12). After this period, centrifuge at 1,400gmax (∼3,900 rpm in an Eppendorf 5415D microfuge using the standard 24-place rotor) for 10 min at 4°C, and then remove and retain the supernatant. 2. Protein Precipitation: It is often necessary to precipitate proteins derived from plant material lysis, to ensure the removal of phytophenols, lipids and other cellular components that may interfere with downstream processing steps. There are several commercial kits available for this purpose; e.g., PlusOne 2-D Clean-Up

Fig. 4. Analysis of chloroplast protein samples by CyDyeDIGE 2D-PAGE. Chloroplasts were isolated from 10-day-old Arabidopsis seedlings using the described protocol, and then analysed using CyDyeDIGE technology as described previously (28,29). An image of a typical pH 3–10 nonlinear 2D-DIGE gel is shown (the pH range is indicated at the top). Two different chloroplast protein samples (one from wild-type Arabidopsis, and another from a mutant) were labeled with minimal CyDyeDIGE fluors, and then analysed simultaneously. The images from the wild-type sample (Cy5) and the mutant sample (Cy3) have been overlaid. If the Cy5 signal were colored red and the Cy3 signal were colored green, spots corresponding to proteins spots corresponding to proteins deficient in the mutant would appear red, whereas proteins enriched in the mutant would appear green. The large (LSU) and small (SSU) subunits of Rubisco are indicated.

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kit (Amersham Biosciences), or PerfectFOCUS (Genotech, St. Louis, Missouri). Alternatively, refer to other chapters in this book. 3. Protein estimation: The protein concentration of the sample(s) in lysis buffer must be determined before proceeding with any proteomics methodologies. There are several commercial kits available for this purpose, but it is imperative that the method used is compatible with samples containing detergents; e.g., RC/DC Protein Assay (Bio-Rad Laboratories Ltd.), or PlusOne 2-D Quant (Amersham Biosciences). 4. Once these various preparative procedures have been completed, the solubilized chloroplast protein samples can be subjected to a variety of different methodologies for proteome analysis, such as difference gel electrophoresis (DIGE) (see Fig. 4).

4. Notes 1. For 10-day-old plants, this is roughly equivalent to ∼6,000–8,000 individuals or ∼20 g tissue. For older plants, few individuals are needed, and the density of plants per plate should be reduced. 2. It is likely that more plates will be needed when working with sick, mutant plants. 3. The method works just as well using a Kinematica PT20 polytron with a small rotor (13 mm diameter) at ∼40% maximum speed, as described previously (30). 4. Prepare 2× CIB and store in 200-mL aliquots in 400 mL Duran bottles at –20°C. Before use, thaw overnight at 4°C, then make CIB by adding 200 mL sterile deionized H2 O and mixing well. 5. Small amounts of 2× CIB are needed for preparation of Percoll gradients, so aliquots of 2× CIB in 50-mL tubes are recommended for storage at –20°C. The buffer can be thawed and frozen several times (up to 5 times). Mix well after thawing. 6. Prepare 50-mL aliquots and store at –20°C. The buffer can be thawed and frozen several times (up to 5 times). Mix well after thawing. 7. Do not wear gloves when handling seeds. The seeds will stick to the gloves because of static electricity. Instead, sterilize hands with IMS and avoid touching the seeds directly. 8. When working with certain, particularly sick mutants, it may be beneficial to use MS medium supplemented with 100 mM (∼3%) sucrose. 9. When using newly thawed aliquots of 2× CIB, mix well before use to obtain a homogeneous solution. 10. Sometimes, yields can be improved by increasing the volume of CIB and increasing the number of rounds of homogenization; this may be particularly advantageous when using large amounts of tissue. 11. When using small rotors (e.g., PTA10S with the Kinematica PT10-35, or the 13-mm rotor with the Kinematica PT20) good homogenization is achieved by moving the small beaker up and down 8–10 times quickly (taking ∼3–4 s in

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total), in each one of the five rounds of homogenization. When working with chlorotic or sick mutants, homogenization times should be reduced according to the severity of the mutant phenotype. For example, when isolating chloroplasts from the ppi1 mutant (28), homogenization time was reduced to ∼1–2 s for each of the five rounds of homogenization. When using larger rotors (e.g., PTA20S with the Kinematica PT10-35), the up and down movement of the beaker is not possible because of the strong suction generated. 12. Buffers containing ASB-14 will solidify at 4°C. Therefore, lysis and solubilization using such buffers should be carried out at room temperature, and the lysis buffer should include a protease inhibitor cocktail (e.g., Complete Protease Inhibitor Cocktail Tablets, Roche Diagnostics Ltd., Lewes, UK).

Acknowledgments We thank Ramesh Patel for technical assistance, and Natalie Allcock and Stefan Hyman (Electron Microscope Laboratory, Faculty of Medicine and Biological Sciences, University of Leicester) for transmission electron microscopy. We are grateful to Sabina Kovacheva and Ramesh Patel for their helpful comments on the manuscript. We thank Marc Boutry (PMA2 H+ -ATPase), Karl-Josef Dietz (PrxII F), Richard Trelease (catalase), Peter Shaw (histone H3), Henrik Scheller (PSI-D), and Kenton Ko (SSU) for generously providing antibodies. This work was supported by the Biotechnology and Biological Sciences Research Council (BBSRC) Genomic Arabidopsis Resource Network (GARNet), by the Royal Society Rosenheim Research Fellowship (to P.J.), and by BBSRC Grants 91/P12928 and BBS/B/03629 (to P.J.).

References 1. Whatley, J. M. (1978) A suggested cycle of plastid developmental interrelationships. New Phytol. 80, 489–502. 2. Lopez-Juez, E. and Pyke, K. A. (2005) Plastids unleashed: their development and their integration in plant development. Int. J. Dev. Biol. 49, 557–77. 3. Leister, D. (2003) Chloroplast research in the genomic age. Trends Genet. 19, 47–56. 4. Nelson, N. and Ben-Shem, A. (2004) The complex architecture of oxygenic photosynthesis. Nat. Rev. Mol. Cell Biol. 5, 971–82. 5. Baginsky, S. and Gruissem, W. (2004) Chloroplast proteomics: potentials and challenges. J. Exp. Bot. 55, 1213–20. 6. Jarvis, P. (2004) Organellar proteomics: chloroplasts in the spotlight. Curr. Biol. 14, R317–R319. 7. van Wijk, K. J. (2004) Plastid proteomics. Plant Physiol. Biochem. 42, 963–77.

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17 Isolation of Plant Cell Wall Proteins Elisabeth Jamet, Georges Boudart, Gisèle Borderies, Stephane Charmont, Claude Laﬁtte, Michel Rossignol, Herve Canut, and Rafael Pont-Lezica

Summary The quality of a proteomic analysis of a cell compartment strongly depends on the reliability of the isolation procedure for the cell compartment of interest. Plant cell walls possess specific drawbacks: (1) the lack of a surrounding membrane may result in the loss of cell wall proteins (CWP) during the isolation procedure; (2) polysaccharide networks of cellulose, hemicelluloses, and pectins form potential traps for contaminants such as intracellular proteins; (3) the presence of proteins interacting in many different ways with the polysaccharide matrix require different procedures to elute them from the cell wall. Three categories of CWP are distinguished: labile proteins that have little or no interactions with cell wall components, weakly bound proteins extractable with salts, and strongly bound proteins. Two alternative protocols are decribed for cell wall proteomics: (1) nondestructive techniques allowing the extraction of labile or weakly bound CWP without damaging the plasma membrane; (2) destructive techniques to isolate cell walls from which weakly or strongly bound CWP can be extracted. These protocols give very low levels of contamination by intracellular proteins. Their application should lead to a realistic view of the cell wall proteome at least for labile and weakly bound CWP extractable by salts.

Key Words: Arabidopsis thaliana; bioinformatics; cell fractionation; cell wall; cell wall protein; plant; proteomics.

1. Introduction Plant cell wall proteins (CWP) present specific complexities in addition to the difficulties usually encountered in proteome analysis, such as protein separation and detection of scarce proteins (1). They are embedded in an insoluble From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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polysaccharide matrix and interact with other cell wall components, making their extraction challenging. Current models of cell wall structure describe the arrangement of their components into two structurally independent and interacting networks, embedded in a pectin matrix (2,3). Cellulose microfibrils and hemicelluloses constitute the first network; the second one is formed by structural proteins. Three types of CWP can be distinguished, according to their interactions with cell wall components (4). CWP can have little or no interactions with cell wall components and thus move freely in the extracellular space. Such proteins can be found in liquid culture media of cell suspensions and seedlings or can be extracted with low ionic strength buffers. We call this fraction “labile proteins,” most of them have acidic pI ranging from 2 to 6 (Fig. 1A). Alternatively, CWP might be weakly bound to the matrix by Van der Waals interactions, hydrogen bonds, and hydrophobic or ionic interactions. Such proteins may be extracted by salts and most of them have basic pI ranging from 8 to 11 (Fig. 1B) so that they are positively charged at the acidic pH of cell walls. Even though most of the cell wall polysaccharides are neutral, negatively

Fig. 1. pIs of labile and weakly-bound CWP. pIs of CWP identified in several proteomic studies (4) were calculated (www.iut-arles.up.univ-mrs.fr/w3bb/d_ abim/compo-p.html) after removal of their predicted signal peptides (http:// psort.nibb.ac.jp/form.html). Three groups of proteins were considered: (A) labile proteins; (B) salt-extracted proteins, i.e. proteins extracted with salt solutions or chelating agent; (C) all proteins. Reprinted from (4), Copyright (2005), with permission from Elsevier.

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charged pectins contain polygalacturonic acid that provides negative charges for interactions with basic proteins. Such interactions would be modulated by pH, degree of pectin esterification, Ca2+ concentration, and by the mobility and diffusion coefficients of these macromolecules (3,5). Finally, CWP can be strongly bound to cell wall components so that they are still resistant to salt-extraction. As examples, extensins are cross-linked by covalent links (6,7) and peroxidases can have a high affinity for Ca2+ -pectate (8). The available techniques described in this chapter allow the extraction of labile and weakly bound CWP. Because labile proteins can be lost during the preparation of cell walls, they must be extracted from tissues by nondestructive techniques such as vacuum infiltration (9), or recovered from liquid culture media from cell suspension cultures or seedlings (10,11). Weakly bound CWP can be extracted with salts or chelating agents from living cells with nondestructive techniques (9,10) or from purified cell walls with destructive techniques. At present, there is no efficient procedure to release CWP strongly bound to the extracellular matrix. Structural proteins, for instance extensins or PRP, can be cross-linked via di-isodityrosine bonds (6,12). Purified cell walls appear as the most suitable material to isolate such proteins. However, until now, extensins have only been eluted with salts before their insolubilization from cell suspension cultures (13). 2. Materials A major problem in proteomics is the occurrence of keratins that can contaminate materials and working solutions. The presence of keratins can prevent the identification of proteins of interest by mass spectrometry. It is necessary to pay attention to all possible sources of contamination at all steps of the following protocols. Powder-free gloves should be permanently worn and washed with soap before their first use. Chemicals should be reserved for proteomic studies and should not be manipulated with spatula. Buffers should be filtered on 0.22 μm pore size filters. Glass plates for electrophoresis should be cleaned with alcohol before use. 2.1. Extraction of Labile or Weakly Bound CWP by Nondestructive Techniques 2.1.1. CWP extraction and Analysis from Liquid Culture Medium of Seedlings 1. Murashige and Skoog (MS) culture medium: Murashige and Skoog (14) liquid medium (Sigma Chemical, St Louis, MO, USA) is supplemented with 10 g/L sucrose and adjusted to pH 5.8 with KOH. 2. PVPP (Sigma, St. Louis, MO, USA) is treated with acid to increase polymerization and to remove metal ions and contaminants. One g PVPP in 10 mL 10%

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Jamet et al. HCl is boiled for 10 min, filtered through a G4 filter, and rinsed until neutral pH is reached. The residue is dehydrated with acetone and grinded in a mortar to obtain a fine powder (15). Low binding 12 kDa cutoff Spectra/Por® cellulose ester (CE) dialysis bags (Merck Eurolab Poly Labo, Strasbourg, France). Centriprep® device (MWCO: 10 kDa) (Millipore Corporation, Bedford, MA, USA). Bradford protein assay (Coomassie® Protein assay Reagent Kit, Pierce, Rockford, IL, USA) (16). Immobilized pH gradient (IPG) buffers and 13 cm-strips pH 4 –7 or 6–11 (GE Healthcare Europe GmbH, Orsay, France). 2-DE (2-dimensional electrophoresis) sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 0.5% (v/v) IPG buffer (pH 4-7 or 6-11), bromophenol blue trace.

2.1.2. CWP Extraction and Analysis from Cell Suspension Cultures 1. Gamborg liquid medium: Gamborg B5 medium supplemented with 20 g/L sucrose, 2.5 μM naphthalene acetic acid (Sigma Chemical, St Louis, MO, USA) and adjusted to pH 5.7 with KOH (17). 2. Cell washing buffer: 50 mM sodium acetate buffer pH 6.5, 10 mM DTT, 1 mM PMSF, 1% ethanol, 50% glycerol. 3. C1 protein extraction buffers: 0.15M NaCl in cell washing buffer. 4. C2 protein extraction buffer: 1M NaCl in cell washing buffer. 5. C3 protein extraction buffer: 0.2M CaCl2 dihydrate in cell washing buffer. 6. C4 protein extraction buffer: 2M LiCl in cell washing buffer. 7. C5 protein extraction buffer: 50 mM 1,2-cyclohexanediamine tetracetic acid (CDTA) in cell washing buffer. 8. Low binding 2 kDa cutoff Spectra/Por® CE dialysis bags (Merck Eurolab Poly Labo, Strasbourg, France). 9. One mL Hi-Trap SP Sepharose cation exchanger (GE Healthcare Europe GmbH, Orsay, France). 10. Hi-Trap SP equilibration buffer: 10 mM MES-KOH, pH 5.2. 11. Hi-Trap SP elution buffer: 10 mM MES-KOH, pH 5.2, 2M NaCl. 12. Econo-Pac® 10DG desalting column (Bio-Rad, Hercules, CA,USA). 13. Desalting column equilibration buffer: 50 mM ammonium formate. 14. Bradford protein assay (Coomassie® Protein assay Reagent Kit, Pierce, Rockford, IL, USA) (16). 15. 1-DE (1-dimensional electrophoresis) sample buffer: 62 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 5% mercapto-ethanol. 16. Resuspending solution: 1M thiourea, 10 mM DTT, 1% (v/v) protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA) in UHQ water (see Note 1). Prepare as required. 17. Immobilized pH gradient (IPG) buffers and 13-cm strips pH 4–7 or 6–11 (GE Healthcare Europe GmbH, Orsay, France).

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18. 2-DE sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 0.5% (v/v) IPG buffer (pH 4–7 or 6–11), bromophenol blue trace.

2.1.3. Extraction and Analysis of CWP from Rosette Leaves 2.1.3.1. Extraction of Proteins 1. Recovering solution: 0.3M mannitol, 66 mM DTT, 330 mM thiourea, 3.3% (v/v) protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA) (see Note 1). Prepare as required. 2. R1 protein extraction buffer: 1M NaCl, in recovering solution. Adjust pH to 6.9 with 0.5N NaOH. Prepare as required 3. R2 protein extraction buffer: 0.2M CaCl2 dihydrate, in recovering solution. Adjust pH to 6.9 with 0.5N NaOH. Prepare as required. 4. R3 protein extraction buffer: 2M LiCl, 0.3 in recovering solution. Adjust pH to 6.9 with 1N NaOH. Prepare as required. 5. R4 protein extraction buffer: 50 mM CDTA, in recovering solution. Adjust pH to 6.9 with 5N NaOH. Prepare as required. 6. Malate dehydrogenase (MDH) assay mixture: 50 mM Tris-HCl, pH 7.8 (2.15 mL), 50 mM MgCl2 hexa-hydrate (300 μL), 150 mM DTT (100 μL), 10 mM NADP (150 μL), 30 mM malic acid (300 μL). Store DTT and NADP solutions in single use aliquots at –20°C. Store malic acid solution at –20°C no longer than one month. Prepare the MDH assay mixture as required 2.1.3.2. Analysis of Labile CWP 1. Low binding 2 kDa cutoff Spectra/Por® CE dialysis bags (Merck Eurolab Poly Labo, Strasbourg, France). Store at –20°C. 2. Resuspending solution: 1M thiourea, 10 mM DTT, protease inhibitor cocktail (1% v/v) in UHQ water. Prepare as required. 3. Immobilized pH gradient (IPG) buffers and 7-cm strips pH 4–7 (GE Healthcare Europe GmbH, Orsay, France). 4. 2 DE-sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 0.5% (v/v) IPG buffer (pH 4–7 or 6–11), bromophenol blue trace. 2.1.3.3. Analysis of Weakly Bound CWP 1. Low binding 2 kDa cutoff Spectra/Por® CE dialysis bags (Merck Eurolab Poly Labo, Strasbourg, France). Store at –20°C. 2. One mL Hi-Trap SP Sepharose cation exchanger (GE Healthcare Europe GmbH, Orsay, France). 3. Hi-Trap SP equilibration buffer: 10 mM MES-KOH, pH 5.2. 4. Hi-trap SP elution buffer: 10 mM MES-KOH, pH 5.2, 2M NaCl. 5. Econo-Pac® 10DG desalting column (Bio-Rad, Hercules, CA, USA). 6. Desalting column equilibration buffer: 50 mM ammonium formate. 7. 1-DE sample buffer: 62 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 5% mercapto-ethanol.

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8. Resuspending solution: 1M thiourea, 10 mM DTT, 1% (v/v) protease inhibitor cocktail in UHQ water. Prepare as required. 9. Immobilized pH gradient (IPG) buffers and 7-cm strips pH 4-7 (GE Healthcare Europe GmbH, Orsay, France). 10. 2 DE-sample buffer: 7M urea, 2M thiourea, 4% (w/v) CHAPS, 65 mM DTE, 0.5% (v/v) IPG buffer (pH 4–7 or 6–11), bromophenol blue trace.

2.2. Extraction of Weakly Bound CWP by Destructive Techniques 2.2.1. Cell wall preparation 1. MS solid medium: Murashige and Skoog (14) liquid medium (Sigma Chemical, St Louis, MO, USA) is supplemented with 20 g/L sucrose and 12 g/L agar, and adjusted to pH 5.8 with KOH. 2. PVPP (Sigma, St. Louis, MO, USA) is treated with acid to increase polymerization and to remove metal ions and contaminants. One g PVPP in 10 mL 10% HCl is boiled for 10 min, filtered through a G4 filter, and rinsed until neutral pH is reached. The residue is dehydrated with acetone and grinded in a mortar to obtain a fine powder (15). 3. Nylon nets (1.5-mm pore size and 25-μm pore size). 4. Waring blender with a 2-L flask (SEB Moulinex, Ecully, France). 5. Grinding buffer: 10 mM acetate buffer, pH 4.6, 0.4M sucrose, 0.2 % (v/v) protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA) (see Note 1). 6. Cell wall purification buffers: 5 mM acetate buffer, pH 4.6, 0.6M or 1M sucrose, 0.2% (v/v) protease inhibitor cocktail. 7. Cell wall washing buffer: 5 mM acetate buffer, pH 4.6.

2.2.2. Extraction and Separation of Proteins 1. H1 protein extraction buffer: 5 mM acetate buffer, pH 4.6, 0.2M CaCl2 , 0.1% protease inhibitor cocktail for plant (Sigma, St. Louis, MO, USA). 2. H2 protein extraction buffer: 5 mM acetate buffer, pH 4.6, 2M LiCl, 0.1% protease inhibitor cocktail. 3. Econo-Pac® 10DG desalting column (Bio-Rad, Hercules, CA,USA). 4. Desalting column equilibration buffer: 50 mM ammonium formate. 5. Bradford protein assay (Coomassie® Protein assay Reagent Kit, Pierce, Rockford, IL, USA) (16). 6. 1-DE sample buffer: 62 mM Tris pH 6.8 (HCl), 2% SDS, 10% glycerol, 5% mercapto-ethanol.

3. Methods The choice of a protocol to extract CWP for proteomic analysis is dependent on the plant material and of the type of proteins to be released from cell walls. Working on living cells is probably the best solution to avoid intracellular

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contamination. This is possible for cell suspension cultures or seedlings grown in liquid medium as well as for any plant organ that can be infiltrated under vacuum with extraction buffers. Both labile and weakly bound CWP can be released. When this is not possible, it is necessary to purify cell walls. The main problem is to avoid intracellular contaminants that will stick nonspecifically to cell walls. Only weakly bound CWP can be extracted from purified cell walls because labile CWP are lost during cell wall preparation. Another important point is the choice of the extraction solution. For example, a solution of 0.3M mannitol infiltrated in living tissues such as leaves can solubilize a few CWP expected to be located only in intercellular spaces. Indeed, identified proteins are acidic, suggesting no ionic interactions with negatively charged cell wall components (9). NaCl is usually used for extraction of proteins retained by ionic interactions in the cell wall. LiCl can extract hydroxyprolinerich glycoproteins from intact cells in Chlamydomonas reinhardii (18). Calcium chloride is probably the most efficient salt to extract CWP (9,19). The ability of acidic and neutral carbohydrates to strongly chelate calcium (20,21) might explain, through a competition mechanism, that proteins or glycoproteins weakly bound to cell wall polysaccharides can be selectively solubilized by CaCl2 . CDTA, a chelating agent, solubilizes Ca2+ -pectate. It releases a small number of proteins having domains of interaction with polysaccharides, notably proteins showing homology to lectins. This suggests an interaction of these proteins with polysaccharides associated to pectins (9). 3.1. Extraction of Labile or Weakly Bound CWP by Nondestructive techniques 3.1.1. Liquid Culture Medium of Seedlings 1. Soak A. thaliana ecotype Columbia seeds (see Note 2) in tap water for 2 h, then sterilize in diluted bleach (2.4% w/v) for 45 min, and rinse several times with deionized water. 2. Sterilized seeds (100 mg) are germinated and grown in MS liquid culture medium in ten 1-L flasks on a rotary shaker (90 rpm) at 26°C in the dark (22). Each flask contains 130 mL of culture medium. After 14 d, etiolated seedlings are harvested and the culture medium is filtered through nylon net (60 μm) to remove cell debris. 3. Collect 900 mL of culture medium (see Note 3). Mix with 9 g PVPP. Shake the mixture at 4°C for at least 30 min, filter and centrifuge to pellet the insoluble residue. Dialyze against 10 L distilled water during 10–12 h at 4°C using a dialysis bag (MWCO: 12 kDa) with three changes. Reduce the volume of the sample by repeated centrifugations (3,500g for 15 min at 4°C) through a Centriprep® device (MWCO: 10 kDa) to about 1 mL. 4. Quantify proteins using the Bradford protein assay.

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5. Dilute 250 μg proteins extracted from culture medium with 250 μL 2-DE sample buffer. Proteins are separated by 2-DE using 13 cm- IPG gel strips pH 4–7 or 6–11 for the first dimension.

3.1.2. Cell Suspension Cultures 1. A cell suspension culture of A. thaliana ecotype Columbia is grown on Gamborg liquid medium. From this culture, 50 mL (25 g) is transferred every 2 wk to 250 mL fresh medium in 1-L Erlenmeyer flask and shaken at 70 rpm in an orbital shaker, under continuous light (30 μE.m−2 .s−1 ) at 22°C. 2. Wash cells of 7-day-old A. thaliana suspension culture with water and pellet them by centrifugation at 200g. Plasmolyze by successive immersion in 25 % glycerol, 50 % glycerol for 10 min each, and finally wash in 50 % cold glycerol. All subsequent extractions are performed at 0°C except otherwise stated. 3. Before protein extraction, wash cells with cell washing buffer to remove contaminant proteins coming from broken cells that nonspecifically stick to cell walls. 4. Extract proteins by washings of the plasmolyzed cells under gentle stirring (30 min) in the proportion of 25 mL of pelleted cells per 50 mL of solution. First extraction is performed with C1 buffer. 5. Wash cells with the same extraction buffer, then with 50 % glycerol before centrifugation at 200g for 5 min. 6. Extract weakly bound CWP with C2, C3, C4, or C5 buffer in the same way (see Note 4). 7. Exhaustively dialyze protein extracts at 4°C against 20 L H2 O using a dialysis bag (MWCO: 2 kDa). Measure the protein content of each extract using the Bradford protein assay. 8. Dialyze against Hi-Trap SP equilibration buffer and apply to a Hi-Trap SP Sepharose column equilibrated with Hi-Trap SP equilibration buffer at 1 mL/min. Elute the retained basic proteins with Hi-Trap SP elution buffer at 1 mL/min. Desalt the basic proteins on an Econo-Pac® 10DG desalting column equilibrated with desalting column equilibration buffer. Freeze-dry the eluate. Resuspend the dry residue in 40 μL 1-DE sample buffer and separate proteins by 1-DE on a 10–17% gradient polyacrylamide gel (16.5 × 13.5 × 0.15 cm). 9. Freeze-dry the acidic and neutral proteins from the Hi-Trap SP Sepharose column effluent. Solubilize the dry residue with a minimal volume of resuspending solution and desalt on an Econo-Pac® 10DG desalting column. Freeze-dry the proteins, dissolve in 2-DE sample buffer and perform a 2-DE using a 7 cm-IPG gel strip pH 4–7 for the first dimension.

3.1.3. Rosette Leaves 3.1.3.1. Extraction of Proteins 1. Sterilize A. thaliana ecotype Columbia seeds by soaking in diluted bleach (2.6% w/v) for 5 min and rinse several times with deionized water. Sow the seeds on

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humid compost in 10 × 10 cm pots and cover the pots with a plastic film. Remove the plastic film after 48 h and transfer the cultures in a growth chamber at 70% humidity, with a photoperiod of 9 h light at 110 μE.m−2 s−1 at 22°C, and 15 h dark at 20°C. Plants should be moderately watered with a nutrient solution once a week. 2. Remove carefully 4- to 5-week-old plants (Fig. 2A) from the pots and wash compost off with deionized water. Cotyledons and yellowish leaves should be systematically removed from plants. Process whole plants for vacuum-infiltration as follows. Make a small noose with a piece of string and pass the root through the noose. Tighten the noose around the collar then twist the root around the string and wrap in parafilm. In a large beaker, immerse completely the rosettes first in distilled water for a few seconds in recovering solution. Put the beaker with the immersed rosettes in a dessicator connected to a vacuum pump (Fig. 2B). Vacuum-infiltrate the rosettes for 2 min after starting the pump. Reintroduce carefully air in the dessicator after vacuum breakage (Fig. 2C). Transfer the infiltrated plants to a centrifuge tube, with the collar at about 1 cm at the edge of the tube (Fig. 2D). Paste the lower part of the root outside of the tube with adhesive tape. Introduce at the bottom of the centrifuge tube 300 μL of recovering solution. Centrifuge infiltrated plants in swinging buckets at 200g for 17 min at

Fig. 2. Vacuum-infiltration of rosette leaves. Four steps of the procedure are illustrated: (A) 4–5 wk-old plants; (B) vacuum-infiltration of immersed rosettes in a dessicator connected to a vacuum pump; (C) rosette leaves after vacuum-infiltration, note the darker part of a leaf after successful infiltration (black arrow); (D) infiltrated plant transferred to a centrifuge tube, note the drop of solution containing the protease inhibitor cocktail at the bottom of the tube (black arrow).

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20°C (see Note 5). Collect the apoplastic washing fluids with a micropipet and estimate the volume. Vacuum-infiltration and centrifugation should be repeated twice. 3. Assay the apoplastic fluids for malate dehydrogenase (MDH) activity to detect cytoplasmic contaminations. Measure MDH activity at room temperature in 3 mL MDH assay mixture and one-twentieth of the volume of the recovered apoplastic fluids. Reduction of NADP is followed at = 340 nm. Pool only those apoplastic washing fluids with no detectable MDH activity. 4. Vacuum-infiltrate rosettes with R1, R2, R3 or R4 buffer. Check for MDH activity on the recovered apoplastic fluids as described above. Discard any apoplastic washing fluids with MDH activity. Pool the remaining apoplastic washing fluids free of MDH activity.

3.1.3.2. Analysis of Labile CWP by 2-DE 1. Exhaustively dialyze the apoplastic washing fluids from rosettes infiltrated with the recovering solution at 4°C against deonized water in low binding 2 kDa cutoff Spectra/Por® CE dialysis bags. Freeze-dry the dialysates. Resuspend the dry residues in 3 mL of resuspending solution and desalt on an Econo-Pac® 10DG desalting column equilibrated with desalting column equilibration buffer for the complete removal of mannitol. Freeze-dry the eluate. 2. Solubilize the dry residue in 2-DE sample buffer and separate proteins by 2-DE using a 7 cm-IPG gel strip pH 4–7 for the first dimension. 3.1.3.3. Analysis of Weakly Bound CWP 1. Exhaustively dialyze the apoplastic washing fluids from rosettes infiltrated with R1, R2, R3, or R4 buffer against Hi-Trap SP equilibration buffer as described above. Apply to a Hi-Trap SP Sepharose column equilibrated with Hi-Trap SP equilibration buffer at 1 mL/min. Elute the retained basic proteins with Hi-Trap SP elution buffer at 1 mL/min. Desalt the basic proteins on an Econo-Pac® 10DG desalting column equilibrated with desalting column equilibration buffer. Freezedry the eluate. Resuspend the dry residue in 40 μL 1-DE sample buffer and separate proteins by 1-DE on a 10–17% gradient SDS-polyacrylamide gel (16.5 × 13.5 × 0.15 cm). 2. Freeze-dry the acidic and neutral proteins in the Hi-Trap SP Sepharose column effluent. Solubilize the dry residue with a minimal volume of resuspending solution and desalt on an Econo-Pac® 10DG desalting column equilibrated with desalting column equilibration buffer. Freeze-dry the proteins and perform a 2-DE.

3.2. Extraction of Weakly Bound CWP by Destructive Techniques 3.2.1. Cell Wall Preparation 1. Soak A. thaliana ecotype Columbia seeds (see Note 2) in tap water for 2 h, then sterilize in diluted bleach (2.4 %) for 45 min, and rinse several times with deionized water. Sow the seeds (150 mg) in a Magenta box (6 × 6 cm) containing

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50 mL of solid MS medium. Grow seedlings at 23°C in the dark for 11 d (see Note 6). Harvest hypocotyls (around 2 cm high) of an average of 20 Magenta boxes as follows. First, remove carefully the solid MS medium carrying the seedlings from each box. Then, cut hypocotyls below cotyledons and above root with a pair of scissors. Wash the 1-cm-long hypocotyls with distilled water onto a nylon net (1.5-mm pore size) to remove all the cut cotyledons and seed coats that stick to hypocotyls (see Note 7). Transfer the hypocotyls into 500 mL of grinding buffer and add PVPP (1g/10g fresh weight of hypocotyls) to complex phenolic compounds. Grind the mixture in cold room using a Waring blender at full speed for 15 min. Separate cell walls from soluble cytoplasmic fluid by centrifugation of the homogenate for 15 min at 1,000g and 4°C. Further purify the pellet by two successive centrifugations in 500 mL of cell wall purification buffers, 0.6M and 1M sucrose respectively. Wash the residue with 3 L of cell wall washing buffer on a nylon net (25-μm pore size) to eliminate all soluble compounds. Grind the resulting cell wall fraction in liquid nitrogen in a mortar with a pestle before lyophilization. Starting with 16 g fresh weight of hypocotyls, this procedure usually results in 1.3 g dry powder.

3.2.2. Extraction of Proteins 1. Typically, 0.65 g of lyophilized cell walls is used for one experiment. Extract proteins by successive salt solutions in this order: two extractions with 6 mL H1 buffer, followed by two extractions with 6 mL H2 buffer. Resuspend cell walls by vortexing for 5-10 min at room temperature, and then centrifuge for 15 min at 4,000g and 4°C. Supernatants from the same extracting buffer are pooled. 2. Desalt supernatants using Econo-Pac® 10DG desalting columns equilibrated with desalting column equilibration buffer. Lyophilize the extracts and resuspend in H 2 O2 . 3. Quantify proteins using the Bradford protein assay. 4. Add 1-DE sample buffer. Separate proteins by 1-DE on a 12.5% SDSpolyacrylamide gel.

3.3. Analysis of CWP 3.3.1. Speciﬁc Constraints for Separation by Electrophoresis and Protein Identiﬁcation by Mass Spectrometry The separation of CWP by classical two-dimensional gel electrophoresis (2-DE) is difficult. Because most CWP are basic glycoproteins (Fig. 1C), they are poorly resolved by this technique (22). They are better separated by 1-DE. However, protocols including chromatographic steps to separate proteins before 1-DE are available (24,25). In this chapter, a method able to separate acidic

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and basic CWP is proposed for a better resolution of these two types of CWP in 2-DE or 1-DE respectively. Frequently, in 1-DE, proteins are not well separated from one another, and a protein sample can contain a mixture of proteins. However, the peptide mass mapping technology using high resolution (< 20 ppm) MALDI-TOF mass spectrometry (MS) permits the identification of several proteins from a mixture. Search engines such as MSFIT from Protein Prospector (http://prospector.ucsf.edu/ucsfhtml4.0/msfit.htm) or MASCOT (http://www.matrixscience.com/search_form_select.html) allow multiple searches. In case of difficulties, proteins can also be identified by peptide sequencing using LC (liquid chromatography)-MS/MS systems (9). 3.3.2. Use of Bioinformatics for the Evaluation of the Efﬁciency of an Extraction Protocol The reliability of protein profiling for a compartment like the cell wall, strongly depends on the quality of the preparation. Unfortunately, the classical methods to check the purity of a particular fraction are not conclusive for proteomic studies, because the sensibility of the analysis by mass spectrometry is 10–1,000 times more sensitive than enzymatic or immunological tests using specific markers. Our experience in the field has shown that the most efficient way to evaluate the quality of a cell wall preparation is (1) to identify all the proteins extracted from the cell wall by mass spectrometry, and (2) to perform extensive bioinformatic analysis to determine if the identified proteins contain a signal peptide, and no retention signals for other cell compartments. Comparison of the results obtained with different programs is necessary to ensure a reliable prediction: PSORT allows predicting any subcellular localization (http://psort.ims.u-tokyo.ac.jp/ form.html); TargetP looks for the presence of signal peptides for protein secretion or of transit peptides for mitochondrion or chloroplast targeting (http://www.cbs.dtu.dk/services/TargetP/); Aramemnon compares the results of several programs predicting the presence of signal peptides and transmembrane domains (http://aramemnon.botanik.uni-koeln.de/). It is then possible to conclude about the quality of the cell wall preparation by calculating the ratio of predicted secreted proteins to intracellular ones. 4. Notes 1. Protease inhibitor cocktail for plant is required to prevent proteolysis during the extraction procedure. Proteolysis induces the production of smaller broken proteins that can be spread over 1-D or 2-D polyacrylamide gels. Thus, proteolysis can prevent the identification of both broken proteins and other proteins of interest

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3. 4.

5.

6.

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by mass spectrometry. Moreover, the occurrence of these polypeptides is a great problem for quantitative and comparative proteomics. Seeds germinate in culture media that are favourable to development of bacteria or fungi. Because of the high amount of seeds (150 mg) introduced in a culture flask or in a Magenta box, the possible contamination events are multiplied. So, seeds should be carefully sterilized, and the healthy state of plants should be very good during their production in greenhouses. Culture media should be processed immediately after recovery. Otherwise, proteolysis may occur even if they are stored frozen. No more than two successive extractions with salt solutions should be performed. Otherwise, cells are damaged and intracellular contaminants are released in the culture medium and can stick non-specifically to cell walls (10). Be careful setting minimal acceleration to avoid seriously damaging the vacuuminfiltrated plants during the centrifugation step. Imperatively centrifuge in swinging buckets to get undamaged plants during spinning. All seedlings should grow at about the same rate to reach the same size after 11 d. If germination is not homogeneous, place the boxes at 6°C during 2 d to allow all seeds to start germination without growth. Then, all boxes can be put at 23°C for 11 d. Cotyledons should be carefully removed. They contain few protein species but each of them in a huge amount. Because of their density, cotyledons co-sediment with cell walls. As a consequence, few cotyledons induce a significant contamination during extraction of cell wall proteins, especially by storage proteins. This contamination prevents the identification of proteins of interest by mass spectrometry.

Acknowledgments The authors are grateful to the Université Paul Sabatier (Toulouse III, France) and the CNRS for support. References 1. Hunter, T. C., Andon, N. L., Koller, A., Yates, J. R. and Haynes, P. A. (2002) The functional proteomics toolbox: methods and applications. J. Chromatogr. B 782, 161–181. 2. Carpita, N. and Gibeaut, D. (1993) Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J. 3, 1–30. 3. Cosgrove, D. J. (2005) Growth of the plant cell wall. Nat. Rev. Mol. Cell. Biol. 6, 850–861. 4. Jamet, E., Canut, H., Boudart, G. and Pont-Lezica, R. F. (2006) Cell wall proteins: a new insight through proteomics. Trends Plant Sci. 11, 33–9. 5. Varner, J. E. and Lin, L.-S. (1989) Plant cell wall architecture. Cell 56, 231–39.

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6. Brady, J. D., Sadler, I. H., and Fry, S.C. (1996) Di-isodityrosine, a novel tetrameric derivative of tyrosine in plant cell wall proteins: a new potential cross-link. J. Biochem. 315, 323–27. 7. Schnabelrauch, L. S., Kieliszewski, M. J., Upham, B. L., Alizedeh, H. and Lamport, D. T. A. (1996) Isolation of pI 4.6 extensin peroxidase from tomato cell suspension cultures and identification of Val-Tyr-Lys as putative intermolecular cross-link site. Plant J. 9, 477–89. 8. Shah, K., Penel, C., Gagnon, J., and Dunand, C. (2004) Purification and identification of a Ca+2 -pectate binding peroxidase from Arabidopsis leaves. Phytochem. 65, 307–12. 9. Boudart, G., Jamet, E., Rossignol, M., et al. (2005) Cell wall proteins in apoplastic fluids of Arabidopsis thaliana rosettes: Identification by mass spectrometry and bioinformatics. Proteomics 5, 212–21. 10. Borderies, G., Jamet, E., Lafitte, C., et al. (2003) Proteomics of loosely bound cell wall proteins of Arabidopsis thaliana cell suspension cultures: a critical analysis. Electrophoresis 24, 3421–32. 11. Charmont, S., Jamet, E., Pont-Lezica, R., and Canut, H. (2005) Proteomic analysis of secreted proteins from Arabidopsis thaliana seedlings: improved recovery following removal of phenolic compounds. Phytochem. 66, 453–61. 12. Held, M. A., Tan, L., Kamyab, A., Hare, M., Shpak, E. and Kieliszewski, M. J. (2004) Di-isodityrosine is the intermolecular cross-link of isodityrosine-rich extensin analogs cross-linked in vitro. J. Biol. Chem. 279, 55474–82. 13. Miller, J. G. and Fry, S. C. (1992) Production and harvesting of ionically wallbound extensin from living cell suspension cultures. Plant Cell Tissue Organ Cult. 31, 61–66. 14. Murashige, T. and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue culture. Physiol. Plant. 15, 473–97. 15. Loomis, W. D. (1974) Overcoming problems of phenolics and quinones in the isolation of plant enzymes and organelles. Meth. Enzymol. 31, 528–45. 16. Ramagli, L. S. and Rodriguez, L. V. (1985) Quantitation of microgram amounts of protein in two-dimensional polyacrylamide electrophoresis sample buffer. Electrophoresis 6, 559–63. 17. Axelos, M., Curie, C., Mazzolini, L., Bardet, C. and Lescure, B. (1992) A protocol for transient gene expression in Arabidopsis thaliana protoplasts isolated from cell suspension cultures. Plant Physiol. Biochem. 30, 123–28. 18. Voigt, J. (1985) Extraction by lithium chloride of hydroxyproline-rich glycoproteins from intact cells of Chlamydomonas reinhardii. Planta 164, 379–89. 19. Smith, J., Muldoon, E., and Lamport, D. (1984) Isolation of extensin precursors by direct elution of intact tomato cell suspension cultures. Phytochem. 23, 1233–39. 20. Angyal, S. (1989) Complexes of metal cations with carbohydrates in solution. Adv. Carbohydr. Chem. Biochem. 47, 1–44. 21. van Buren, J. (1991) Function of pectin in plant tissue structure and firmness in, The Chemistry and Technology of Pectin (Walter, R. H. ed.), Academic Press, New York, pp. 1–22.

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22. Bardy, N., Carrasco, A., Galaud, J. P., Pont-Lezica, R. and Canut, H. (1998) Free-flow electrophoresis for fractionation of Arabidopsis thaliana membranes. Electrophoresis 19, 1145–53. 23. Rabilloud, T. (2002) Two-dimensional gel electrophoresis in proteomics: old, old fashioned, but still climbs up the mountains. Proteomics 2, 3–10. 24. Stasyk, T. and Huber, L. A. (2004) Zooming in fractionation strategies in proteomics. Proteomics 4, 3704–16. 25. Lescuyer, P., Hochstrasser, D. F., Sanchez, J. C. (2004) Comprehensive proteome analysis by chromatographic protein prefractionation. Electrophoresis 25, 1125–1135.

18 Isolation and Fractionation of the Endoplasmic Reticulum from Castor Bean (Ricinus communis) Endosperm for Proteomic Analyses William J. Simon, Daniel J. Maltman, and Antoni R. Slabas

Summary This chapter describes the preparation and isolation of highly purified endoplasmic reticulum (ER) from the endosperm of developing and germinating castor bean (Ricinus communis) seeds to provide a purified organelle fraction for differential proteomic analyses. The method uses a two-step ultracentrifugation protocol first described by Coughlan (1) and uses sucrose density gradients and a sucrose flotation step to yield purified ER devoid of other contaminating endomembrane material. Using a combination of one dimensional (1D) and two dimensional (2D) gel electrophoresis the complexity and reproducibility of the protein profile of the purified organelle is evaluated prior to detailed proteomic analyses using mass spectrometry based techniques.

Key Words: Castor bean; electrophoresis; endoplasmic reticulum; proteomics; Ricinus communis.

1. Introduction The endoplasmic reticulum (ER) is a specialized endomembrane system within all cells responsible for a number of important biological processes including, protein folding, protein sorting, and secretion (2), and protein Nglycosylation (3). In the seeds of higher plants the ER is the processing site for the synthesis of storage proteins (4) and is also the site for fatty acid modification (5–7), triacyglycerol (TAG) biosynthesis (8), complex lipid biosynthesis and the primary site for membrane biogenesis. From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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Understanding the processes of TAG biosynthesis and complex lipid partitioning, particularly in the major commercially grown oil producing plant species, is an important step towards maximizing both oil quality and yield from these crops. This is particularly so if advances in molecular biology are to be used to rationally design new crops using transgenic technologies. Although the plant ER plays a central role in metabolism there have been a limited number of studies on the organelle due to the difficulties associated with its isolation and purification. In order to overcome this we have taken advantage of the liquid properties of castor bean endosperm to prepare preparative amounts of ER using modifications of the method first described by Beevers (9). The ER of developing seeds is the major site of synthesis of both lipid and storage proteins – such biosynthetic capacity is lost during germination. In order to elucidate the changes in the ER during seed development and germination we have prepared material from both to allow for a detailed differential proteomic analysis of the two tissues (10). 2. Materials 2.1. Tissue Homogenization and ER Puriﬁcation 1. 5% Hypochlorite solution in water (see Note 1) for seed surface sterilization. 2. Homogenization medium: 500 mM sucrose, 10 mM KCl, 1 mM EDTA, 1 mM MgCl2 2 mM dithiothreitol (DTT), 0.1 mM phenylmethyl-sulfonyl fluoride (PMSF), 150 mM Tricine-KOH pH 7.5. Keep at 4°C on ice. 3. Sucrose solutions for gradient formation: prepare fresh in sterile water 50 mL of each of the following, 20%, 30%, 40%, and 60% sucrose each containing 1 mM EDTA and 0.1 mM PMSF. Keep at 4°C on ice. 4. 10% glycerol solution for –80°C storage of purified ER fractions. 5. The following Beckman centrifuges, rotors and centrifuge tubes: Avanti 30 centrifuge, L-70, and Optima TLX ultra-centrifuges, F0850, SW28, SW41, and TL100.4 rotors, 326823 and 344061 ultra-clear tubes and 343776 micro tubes.

2.2. Protein Estimations 1. Protein standard: Bovine serum albumin (BSA) 1.0 mg/mL in water. 2. Protein assay kit (modified Bradford) from BIO-RAD.

2.3. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis of Puriﬁed ER The SDS-PAGE method described is a modification of that originally described by Laemmli (11). All reagents are from BIO-RAD.

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1. SDS-PAGE resolving gel (10%): 3.33 mL Acrylamide-Bis solution (37.5:1), 2.5 mL 1.5M Tris-HCl pH 8.8, 100 μL of 10% SDS, 100 μL of 10% ammonium persulphate, 10 μL of TEMED in a final volume of 10 mL water. 2. SDS-PAGE stacking gel (4%): 1.3 mL Acrylamide-Bis solution (37.5:1), 2.5 mL 0.5M Tris-HCl pH 6.8, 100 μL of 10% SDS, 100 μL of 10% ammonium persulphate, 10 μL of TEMED (added last once all other constituents are mixed) in a final volume of 10 mL water. 3. Water saturated isobutanol solution: Mix together equal volumes of water and isobutanol in a glass bottle and shake vigorously to mix. Allow to settle and use the top layer. 4. Gel running buffer: 25 mM Tris-HCl pH 8.3, 250 mM glycine, 0.1% SDS (see Note 2). 5. Sample loading buffer: 50 mM Tris-HCl pH 6.8, 100mM DTT, 2% SDS, 0.1% bromophenol blue, 10% glycerol (see Note 3). 6. Dalton Mark VII molecular weight markers (Sigma) made up in sample loading buffer as the manufacturers instructions.

2.4. Mass Spectrometer Compatible Silver Staining Prepare all reagents fresh immediately before use. 1. Fixing solution: 10% acetic acid, 40% methanol. 2. Sensitizing solution: 75 mL methanol, 10 mL sodium-thiosulphate (5 %), 17 g sodium-acetate in 250 mL water. 3. Washing solution: water. 4. Silver solution: 0.25% silver nitrate in water. 5. Developing solution: 6.25 g sodium carbonate, 100 μL formaldehyde in 250mL water. 6. Stop solution: 3.65 g ethylenediaminetetracetic acid (EDTA) in 250 mL water.

2.5. Mini 2 Dimensional Gels (2D gels) 1. The method described in this text uses the Multiphor II ™ electrophoresis unit combined with the Immobiline Dry Strip™ (IPG) kit and the Immobiline Dry Strip Reswelling Tray™ from GE Healthcare for the first dimension isoelectric focusing (IEF) step of 2D electrophoresis. The method should be adaptable to IEF kit from any manufacturer that uses IPG strips, however focusing parameters may need to optimized. For a useful guide to 2D electrophoresis see (12). 2. Immobiline Dry Strip 7 cm ready made IPG strips pH 3–10 from GE Health care. 3. Lysis-rehydration buffer: 9M urea, 2M thiourea, 4% CHAPS (3-[(3 cholamidopropyl)dimethylammonio]-1-propanesulfonate), 1% DTT, 2% carrier ampholytes pH 3–10 (GE Healthcare). Buffer is made up without DTT and carrier ampholytes, aliquoted and stored at –80°C. Once thawed DTT and carrier ampholytes are added immediately before use.

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4. Equilibration buffer 1: 50 mM Tris-HCl pH 8.8, 6M urea, 30% glycerol, 10% SDS, 1% DTT. Prepare 10mL. 5. Equilibration buffer 2: 50 mM Tris-HCl pH 8.8, 6M urea, 30% glycerol, 10% SDS, 4.8% iodoacetamide, 0.01% bromophenol blue. Prepare 10 mL. 6. Low melting point (LMP) agarose solution: 5% made up in SDS-PAGE running buffer. Prepare 20 mL for each mini 2D gels being run by heating the solution to 100°C immediately before use. 7. SDS-PAGE gel reagents as described in Section 2.3. 8. Mass spectrometer compatible silver staining reagents as described in Section 2.4.

3. Methods 3.1. Isolation of ER Membranes from Germinating and Developing Castor Bean Endosperm 1. In order to make a proteomic comparison between the ER from germinating and developing castor bean, material must be prepared from the seeds at both growth stages. For the germinating sample mature seeds are first surface sterilized for 1.0 min in 5% hypochlorite solution and then washed overnight in running tap water. The soaked seeds are then sown in moist vermiculite and germinated in the dark at 30°C for 3–4 d before dissecting out the endosperm. For the developing material seedpods are harvested from castor plants 25 d after flowering (see Note 4) and individual seeds are removed for processing. 2. Take 40–50 seeds (see Note 5) from each of the germinating or developing samples and carefully dissect out the endosperm and remove the embryo and cotyledons. Place the endosperm halves into a large glass Petri dish containing 10 mL ice-cold homogenization buffer and manually chop them for 10 min on ice using two single-sided razor blades (see Note 6). Carry out all further procedures at 4°C unless otherwise stated. 3. Filter the crude homogenate through a 100-μm nylon mesh to remove large debris and transfer the filtrate to 50-mL ultra-clear centrifuge tubes and centrifuge in a Avanti 30 bench-top centrifuge using a F0850 rotor at 1,000g at 4°C for 15 min. 4. During this centrifugation step prepare in two Beckman Ultraclear 25 × 89-mm centrifuge tubes, discontinuous sucrose density gradients consisting of 7 mL of 20% sucrose carefully layered on top of 13 mL of 30% sucrose. Both sucrose solutions contain 1 mM EDTA and 0.1 mM PMSF. 5. With a glass rod carefully remove and discard the fat pad (see Note 7) which has formed on top of the supernatant during centrifugation of the homogenate and divide the supernatant into two halves each of which is carefully layered onto the top of the prepared sucrose gradients (step 4 above). 6. Transfer the tubes to the buckets of the Beckman SW28 rotor and centrifuge at 100,000g for 2 h at 2°C in a Beckman L70 ultra-centrifuge. 7. Following centrifugation mount the tubes vertically in a laboratory clamp positioned under a lamp and the membranes should be clearly visible as a distinct band at the 20-30% sucrose interface (Fig. 1). Without disturbing the

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Fig. 1. Following centrifugation at 100,000g for 2 h the centrifuge tube is carefully removed from the rotor without disturbing the sucrose gradient and mounted in a laboratory clamp positioned under a lamp. This enables the membrane layer to be clearly seen (marked by the arrow) at the 20–30% sucrose interface (∼1.12 g/cm3 density). As shown a hypodermic needle is used to pierce the centrifuge tube wall just below the membrane layer and the membranes are removed using a syringe.

sucrose gradient carefully pierce the wall of the tube with a hypodermic needle immediately below the interface and using a syringe remove the membrane layer (Fig. 1). 8. Pool the membrane fractions from both tubes and mix with an equal volume of ice cold 60% sucrose containing 1 mM EDTA and 0.1 mM PMSF. 9. Pipet 4.0 mL of diluted membrane fraction into the bottom of Beckman Ultraclear 14 × 89 mm centrifuge tubes and carefully overlay with 3 mL of 40% sucrose solution, 3 mL of 30% sucrose solution and 2 mL of 20% sucrose all containing 1 mM EDTA and 0.1 mM PMSF.

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10. Transfer the tubes to the buckets of the Beckman SW41 rotor and centrifuge at 250,000g for 22 h at 2°C in a Beckman L70 ultra-centrifuge. During this centrifugation step the ER fraction will float through the dense sucrose layers to resolve as a highly purified fraction at the 20–30% sucrose interface (∼1.12 g/cm3 density. 11. Carefully remove the purified ER from each tube as described in step 7 and pool together. Dilute with an equal volume of ice-cold water and aliquot into Beckman thick walled 500-μL polycarbonate centrifuge tubes. 12. Transfer the tubes to the Beckman TL100.4 rotor and pellet the membranes by centrifugation in a Beckman Optima TLX ultracentrifuge at 250,000g for 45 min.

Pour off the supernatant and re-suspend the membrane pellet in a minimum volume of 10% (v/v) glycerol aliquot into tubes suitable for storage, snap freeze in liquid nitrogen and store at –80°C (see Note 8). 3.2. Protein Estimation – Modiﬁed Bradford 1. Using the 1.0 mg/mL BSA stock solution (Section 2.2.1) prepare a calibration curve as outlined in Table 1. Vortex solutions and incubate at room temperature for 15 min. 2. In duplicate mix 2 μL of sample with 798 μL of water and 200 μL of Bradford reagent. Vortex solutions and incubate at room temperature for 15 min. 3. Using a spectrophotometer measure the absorbance of the standards and samples at 595 nm.

Plot the calibration curve and estimate the protein concentration of the samples from this curve. 3.3. (SDS-PAGE) Analysis of Puriﬁed ER from Developing and Germinating Castor Bean Endosperm 1. For SDS-PAGE analyses we use the mini Protean gel kit from BIO-RAD although any gel format may be used including commercially available precast gels. Table 1 Bradford assay calibration curve standard dilutions. BSA Series (μg) 1mg/mLBSA (μL) H2 O (μL) Bradford reagent (μL)

0

1

2

5

10

15

20

40

800 200

1 799 200

2 798 200

5 795 200

10 790 200

15 785 200

20 780 200

40 760 200

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2. Assemble the gel kit with spacers for 0.75 mm gels and prepare 10 mL of resolving gel solution as outlined in Section 2.3.1. Using a Pasteur pipet carefully pipet the gel solution between the plates avoiding air bubbles. Leave sufficient space between the top of the gel solution and the top of the glass plates to allow for the gel comb and approximately 1 cm of stacker gel. 3. Overlay the gel solution with water-saturated isobutanol and allow the gel to polymerize at room temperature which should take around 30 min. 4. Pour off the isobutanol and rinse the top of the gel thoroughly with several washes of water. 5. Prepare 10 mL of stacking gel solution as described in Section 2.3.2 and using a Pasteur pipet carefully overlay this gel solution onto the top of the polymerized resolving gel. Insert the gel comb avoiding the formation of air bubbles and allow the stacking gel to polymerize, which should take around 30 min at room temperature. 6. While the stacking gel is polymerizing prepare 800 mL of 1× gel running buffer as described in Section 2.3.4. 7. Once the stacking gel has set carefully remove the gel comb and rinse out the wells with running buffer. Assemble the gel kit and fill the upper and lower buffer compartments with running buffer. 8. For SDS-PAGE analyses of purified ER fractions using mass spectrometer compatible silver stain dissolve an aliquot of the purified preparation equivalent to between 10 and 20 μg of total protein in SDS-PAGE sample loading buffer (see Note 9), centrifuge for 5 min in a bench-top microfuge at maximum speed and load directly into the washed wells of the gel. 9. Include at least one lane of SDS-VII molecular weight markers as described in Section 2.3.6. on every gel. 10. Connect the electrophoresis tank to the power supply and run the electrophoresis at 100 volts until the bromophenol blue dye front passes through the stacker gel into the resolving gel. At this point increase the voltage to 200 volts and continue until the dye front reaches the end of the gel. 11. Switch off the power supply, disassemble the gel kit and carefully transfer the gel to a clean polythene container for staining.

3.4. Mass Spectrometer Compatible Silver Staining 1. Mass spectrometric methods for proteomic analyses are highly sensitive techniques capable of detecting and identifying subfemtomole amounts of protein in a sample. For this reason contamination of samples particularly with human keratin can be a major problem during analyses. To avoid this as much as possible use freshly prepared staining reagents, handle gels as little as possible and use clean containers for all staining procedures. 2. Following electrophoresis transfer the gel to a clean staining tray and fix the proteins in the gel by incubating the gel in 100 mL of fixing (Section 2.4.1) solution (see Note 10) for 15 min with gentle shaking (see Note 11) at room temperature. Repeat this step with a second 100 mL of fixing solution.

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3. Discard the fixing solution and add 100 mL of sensitizing solution (Section 2.4.2). Incubate the gel for 30 min at room temperature with gentle shaking. 4. Discard the sensitizing solution and wash the gel thoroughly with 3 × 5 min washes with fresh water. 5. Pour off the last water wash, add 100 mL of silver solution (Section 2.4.4) and incubate the gel for 20 min at room temperature with gentle shaking. 6. Discard the silver solution and wash the gel thoroughly with 3 × 1 min washes with fresh water. 7. Pour off the last water wash, add 100 mL of developing solution (Section 2.4.5) and incubate the gel for 4 minutes at room temperature with gentle shaking. Monitor the intensity of the staining during this incubation period and if necessary lengthen or shorten the developing time to achieve bands of reasonable intensity without staining the gel background. 8. Discard the developing solution, add 100 mL of stopping solution (Section 2.4.6) and incubate the gel for 10 min at room temperature with gentle shaking. 9. Discard the stopping solution and wash the gel thoroughly with 3 × 5 min washes with fresh water. This staining procedure should result in discrete protein bands stained from brown to black, depending on protein concentration, against a clear gel background. A representative 1D-SDS PAGE gel showing three independent Developing ER S

1

2

Germinating ER 3

1

2

3

kDa 66 45 36 29 24 20 14

Fig. 2. Protein estimations were made on each preparation using the modified Bradford procedure as described in Section 3.2. Protein samples (10 μg) were mixed with SDS-PAGE sample buffer vortexed at room temperature for 10 min and microfuged at maximum speed before loading into wells. Electrophoresis was carried out as described in Section 18.3.3. Lane S contains molecular weight markers. The protein profiles are highly reproducible between preparations and clear differences are seen between the developing and germinating ER samples on these gels.

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castor bean ER preparations from both developing and germinating endosperm stained using this protocol is shown in Fig. 2.

3.5. Mini 2D Gel Protein Proﬁling of Puriﬁed ER from Developing and Germinating Castor Bean Endosperm 1. For differential proteomic analyses of the developing and germinating ER a number of independent biological samples need to be prepared. This is to allow for the true biological differences between the two tissue types to be compared and to enable the elimination of technical artifacts such as gel to gel variability, sample solubilization and staining artifacts. The use of mini 2D gels allows the rapid and detailed evaluation of the reproducibility and complexity of the protein profiles of the purified ER fraction from independent preparations before beginning large-scale proteomic profiling experiments. Fig. 3 shows the mini 2D gel profiles from three independent biological preparations of germinating and developing ER prepared using the protocols outlined in this text. 2. To an aliquot of purified ER material equivalent to 50 μg of total protein from each preparation add 4 volumes of ice-cold absolute acetone and incubate on ice for 60 min. Pellet the proteins from this 80% acetone precipitation step by centrifugation for 10 min in a microfuge at maximum speed. 3. Allow the pellet to air dry but do not over-dry. 4. Add 125 μL (see Note 12) of 2D lysis-rehydration buffer (Section 2.5.1) to the pellet and gently disperse it using a pipetman fitted with a 100 μL pipet tip. Sonicate the sample for 10 min in a sonicating water bath and incubate on a vortex shaker at 30°C for 60 min. Centrifuge for 10 min in a microfuge at maximum speed. 5. Using the reswelling tray rehydrate an IPG strip with the entire solubilized sample for a minimum of 6 h but ideally overnight (see Note 13). 6. Assemble the Multiphor kit for isolelectric focusing. 7. Using forceps remove the re-hydrated IPG strip from the reswelling tray, rinse with water and gently blot dry. 8. Place the rehydrated IPG strip in the plastic insert of the IEF tray, position the electrodes and connect them to the power supply. 9. Fill the IEF tray with mineral oil so that the gel strip is completely covered. 10. Fit the lid to the Multiphor unit and begin isoelectric focusing (IEF). 11. IEF is typically carried out in three phases or voltage steps (see Note 14). For 7-cm IPG strips with a sample loading of 50 μg of purified ER fraction these steps are outlined in Table 2. 12. While the IEF gel is running prepare an SDS-PAGE mini gel for the second dimension. For this gel no stacking gel is required and the gel should be prepared with 1-mm spacers to allow for the IPG strip to be positioned on the gel surface. 13. Prepare 10 mL of resolving gel solution as described in Section 2.3.1. Using a Pasteur pipet carefully pipette the gel solution between the gel plates avoiding air bubbles until the solution is about 0.8 cm from the top of the lower glass plate.

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Developing pH 10 S

pH 3.0

pH 10 S

Fig. 3. ER was independently purified from developing and germinating castor bean in three separate experiments using the protocols described in the text. Molecular weight standards were loaded adjacent to the basic end of the IEF strip and are marked S. An aliquot equivalent to 50 μ g of total protein from each preparation was evaluated on mini 2D gels (pH 3.0–10) and stained with mass spectrometer compatible silver stain. As can be clearly seen the independent preparations are reproducible and there are significant differences in the proteomic profiles between the two tissues.

14. Overlay the gel solution with water-saturated isobutanol and allow the gel to polymerize at room temperature until the IEF is completed. 15. Following IEF disconnect the gel kit from the power supply and disassemble the unit (see Note 15). 16. Place the focused IPG strip into equilibration buffer 1 (Section 2.5.4) and incubate at room temperature for 15 min with gentle shaking. 17. Transfer strip to equilibration buffer 2 (Section 2.5.5) and incubate at room temperature for 15 min with gentle shaking.

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Table 2 Isoelectric focusing parametersa step

volts

mA

Watts

Time (h)

kVh

1 2 3

200 3500 3500

2 2 2

5 5 5

0.01 1.30 1.00

2.8 2.2

a Step 1 is a low voltage step which minimizes sample aggregation. Voltage is then gradually increased during step 2 to the final focusing voltage and held at this voltage during step 3 until focusing is complete.

18. During this second equilibration step pour of the isobutanol solution from the polymerized second dimension gel, wash the gel surface extensively with water and blot dry. 19. Prepare low melting point agarose solution as described in Section 2.5.6. 20. Remove the gel strip from the equilibration solution and place it resting on its edge on a piece of moistened filter paper to allow any excess equilibration solution to drain away. 21. Using forceps position the strip between the glass plates of the second dimension gel with its backing strip against one of the glass plates and gently push the IPG strip down until the entire lower edge is in contact with the SDS-PAGE gel surface. Ensure there are no air bubbles trapped between the IPG strip and the gel. 22. If required SDS-VII molecular weight markers can be spotted onto a small piece of filter paper and positioned in contact with the SDS-PAGE gel surface at one end of the IPG strip. 23. Overlay the IPG strip and with low melting point agarose solution that has cooled to between 40–50°C. 24. Assemble the gel kit, connect it to the power supply and begin running the 2nd dimension electrophoresis at 60 volts for 10 min. This will allow the proteins to migrate from the IPG strip into the SDS-PAGE gel. Increase the voltage to 150 volts and continue the electrophoresis until the bromophenol dye front has reached the end of the gel. 25. Following electrophoresis stain the gels using the mass spectrometer compatible silver protocol described in Section 3.4.

4. Notes 1. Unless otherwise stated “Water” in this text refers to high purity water with a resistivity of 18 M-cm. 2. SDS-PAGE running buffer can be made up as a 10× stock solution, stored at room temperature and diluted as required for use.

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3. SDS-PAGE sample loading buffer can be conveniently made up for use as a 5× stock solution, divided into aliquots and stored at –20°C. 4. In order to ensure that developing seeds are collected at the correct developmental stage plants are monitored daily during growth and flowers are tagged as they emerge. The seedpods are then allowed to develop and are harvested after 25 d. 5. The protocol described here has been optimised to yield the maximum amount of purified ER devoid of contamination with other plant endomembrane material. Our attempts to scale up this procedure using higher seed numbers as starting material have only resulted in contaminated preparations. We recommend that for scale up use the protocol as described but increase the number of preparations. 6. Carefully chopping the endosperm with single sided razor blades allows for successful tissue homogenisation without significant membrane disruption. 7. During the low speed centrifugation step of the crude homogenate to remove cell debris a fat pad will form on top of the supernatant in the centrifuge tube. It is important that this fat pad is carefully removed without disruption before the supernatant containing the membrane fraction is loaded onto the sucrose gradient. 8. The typical yield of purified ER from a Ricinus germinating or developing preparation is approx 1.0 mg. We usually re-suspend the final pellet of purified ER in 500 μL of 10% (v/v) glycerol and divide into 5× 100-μL aliquots for storage at –80°C. 9. Samples for SDS-PAGE analyses are best prepared by adding a volume of 5 x sample buffer to an aliquot of sample equivalent to 10–20 μg of total protein and then diluting to 5 times the volume to give a sample ready for loading in 1× sample buffer. 10. For mass spectrometric compatible silver staining use 100mL of each solution for each mini gel being processed. 11. Gels can be left in fixing solution overnight or longer if necessary until staining is required. During staining procedures gels are shaken gently on a gel rocker platform or on an orbital shaker set at low speed. 12. The volume of 2D lysis-rehydration buffer required to solubilize a sample is dependent on the dimensions of the IPG strip being re-hydrated. For mini gels using 7-cm IPG strips the volume is 125 μL, for 11-cm IPG strips 200 μL, for 13-cm IPG strips 250 μL and for 18-cm large format gels 350 μL. 13. Using the IEF reswelling tray up to 12 IPG can be rehydrated with samples at the same time. 14. The voltage and duration of IEF is dependent on the pH and length of the IPG strip being used and on the total amount of protein loaded onto the gel. For guidelines on the parameters used with different IPG strips see the technical data sheets supplied with the IPG strips by the manufacturers. 15. Following IEF gel strips can be equilibrated immediately ready for the SDSPAGE second dimension step or they can be stored at –20°C and the equilibration and second dimension done at a later date.

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References 1. Coughlan, S., Hastings, C., and Winfrey Jnr, R. J. (1996) Molecular charcterisation of plant endoplasmic reticulum – identification of protein disulphide-isomerase as the major reticuloplasmin. Eur. J. Biochem 235, 215–24. 2. Vitale, A. and Denecke, J. (1999) The endoplasmic reticulum – gateway of the secretory pathway. The Plant Cell 11, 614–28. 3. Helenius, A. and Aebi, M. (2004) The role of N-linked glycans in the endoplasmic reticulum. Ann. Rev. Biochem, 73, 1019–49. 4. Galil, G., Sengupta-Gopalan, C., and Ceriotti, A. (1998) The endoplasmic reticulum and its role in protein maturation and biogenesis of oil bodies. Plant. Mol. Biol 38, 1–29. 5. Cassagne, C., Lessire R., Bessoule, J., et al. (1994) Biosynthesis of very long chain fatty acids in plants. Prog. Lipid Res. 33, 55–69. 6. Van de Loo, F.J., Broun, P., Turner, S., and Somerville, C. (1995) Expressed sequence tags from developing castor seeds. Proc. Natl. Acad. Sci. USA. 92, 6743–47. 7. Shanklin, J. and Cahoon, E. B. (1998) Desaturation and related modifications of fatty acids. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49, 611–41. 8. Kennedy, E. P. (1961) Biosynthesis of complex lipids. Fed. Proc. Fed. Am. Soc. Exp. Biol. 20, 934–40. 9. Beevers, H. (1979) Microbodies in higher plants. Annu. Rev. Plant Physiol. 30, 159–73. 10. Maltman, D. J., Simon, W. J., Wheeler, C. H., Dunn, M. J., Wait, R., and Slabas, A. R. (2002) Proteomic analysis of the endoplasmic reticulum from developing and germinating seed of castor (Ricinus communis). Electrophoresis 23, 626–39. 11. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–85. 12. Berkelman, T. and Stenstedt, T. (ed.) (1998) 2-D Electrophoresis using immobilized pH gradients – principles and methods. Amersham Pharmacia Biotech handbook.

19 Cell Wall Fractionation for Yeast and Fungal Proteomics Aida Pitarch, César Nombela, and Concha Gil

Summary The cell wall is an external envelope shared by yeasts and filamentous fungi that defines the interface between the microorganism and its environment. It is an extremely complex structure consisting of an elastic framework of microfibrillar polysaccharides (glucans and chitin) that surrounds the plasma membrane and to which a wide array of different proteins, often heavily glycosylated, are anchored in various ways. Intriguingly, these cell wall proteins (CWPs) play a key role in morphogenesis, adhesion, pathogenicity, antigenicity, and as a promising target for antifungal drug design. However, the CWPs are difficult to analyze because of their low abundance, low solubility, hydrophobic nature, extensive glycosylation, covalent attachment to the wall polysaccharide skeleton, and high heterogeneity. We describe a typical procedure of cell wall fractionation to isolate and solubilize different CWP species from yeasts and filamentous fungi according to the type of linkages that they establish with other wall components and under suitable conditions for following reproducible proteomic analyses. CWPs retained noncovalently or by disulfide bonds are first extracted from isolated yeast or fungal cell walls by detergents and reducing agents. Subsequently, CWPs covalently linked to or closely entrapped within the internal glucan-chitin network are sequentially released either by mild alkali conditions or by enzymatic treatments first with glucanases and then with chitinases. This strategy is a powerful tool not only for obtaining an overview of the sophisticated cell wall proteome of yeasts and filamentous fungi, but also for characterizing mechanisms of incorporation, assembly and retention of CWPs into this intricate cellular compartment and their interactions with structural wall polysaccharides.

Key Words: Cell wall; cell wall proteins; fractionation; fungus; GPI proteins; PIR proteins; proteomics; yeast.

From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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Fig. 1. Schematic representations of the cell wall from S. cerevisiae and C. albicans, and molecular modules of CWPs solubilized by the present procedure of cell wall fractionation. The cell wall of S. cerevisiae and C. albicans basically consists of -1,3 and -1,6-glucans, chitin, mannoproteins and proteins. -1,3-glucan and chitin form an elastic microfibrillar polysaccharide skeleton surrounding the plasma membrane to which mannoproteins are attached through -1,6-glucan, alkali-sensitive bonds, and/or other hitherto uncharacterized linkages. Cell wall proteins (CWPs; mannoproteins and

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1. Introduction The cell wall is an intricate structure common to yeasts and filamentous fungi that surrounds the plasma membrane and is strategically placed at the interface between the cell and its environment, including the host (1–3). This external envelope, accountable for 20–30% of the cell dry weight, is essential for the survival of the microorganism. In fact, it is involved in many vital functions, such as physical protection, osmotic stability, selective permeability barrier, immobilized enzyme support, cell-cell interactions (e.g., cell recognition and adhesion) and morphogenesis, to name but a few. In pathogen fungi, this cellular compartment further takes an active part in virulence, pathogenicity, antigenicity, immunomodulation of the immune response, and adhesion to host substrates (2). Most importantly, its essential nature and its fungal specificity (given its absence in mammalian cells) interestingly make the cell wall an attractive target site (see Note 1) to design antifungal drugs with selective toxicity for human pathogen fungi, such as Candida albicans or Aspergillus fumigatus, among others (4–6). Although the cell wall structure and organization have been investigated most extensively in the prototype yeast Saccharomyces cerevisiae, a similar molecular model is also applicable for other ascomycetes, and in particular for C. albicans, a dimorphic fungus capable of growing either in yeast form or as hyphae (see Note 2) (1,7,8). The cell wall of S. cerevisiae is mainly composed of glucans (with -1,3 and -1,6 linkages), chitin (N-acetylglucosamine polymers), and proteins (often highly O- and/or N-mannosylated) interconnected by covalent and/or non-covalent bonds, leading to an elevated complexity (see Fig. 1). -1,3-glucan, the major component of the cell-wall (electron-transparent) inner layer, forms an elastic three-dimensional microfibrillar framework, encircling the cell, to which other wall constituents are covalently anchored. Chitin is often cross-linked to the -1,3-glucan microfibrillar backbone on its inner side (close to the plasma membrane) and, to a lesser extent, to short sidechains of -1,6-glucan. This N-acetylglucosamine polymer presents low levels (see Note 2), except in the budding neck ring, in the primary septum, in and around the bud scars, or under stress conditions. Both -1,3-glucan and chitin (structural wall polysaccharides) provide mechanical strength and elasticity Fig. 1. proteins) can also be loosely associated, either by non-covalent bonds or through disulfide bridges, with other covalently linked CWPs. See Introduction for further information. The callouts depict details for potential mechanisms of CWP retention into the cell wall on the basis of the procedure of cell wall fractionation described in this chapter to isolate and solubilize different CWP species from yeasts and filamentous fungi.

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to the cell wall. -1,6-glucan, a flexible minor wall component, interconnects certain cell wall proteins (CWPs), the so-called glycosyl phosphatidylinositol (GPI)-CWPs, with -1,3-glucan (∼90% of GPI-CWPs) or chitin (∼10% of GPI-CWPs) through a phosphodiester bridge in their GPI remnant (see Note 3). The CWPs are mostly located on the outside of this -1,3-glucanchitin network (i.e., at the cell-wall electron-dense outer layer) and, in minor amounts, throughout the cell wall, determining its porosity. These CWPs can be: 1. Loosely associated, either noncovalently or through disulfide bonds, with other cell wall components. This group of CWPs comprises (1) soluble precursor forms of covalently linked CWPs, (2) proteins related to the biosynthesis and modulation of wall constituents, such as -1,3-glucosyltransferase (Bgl2p), exoglucanase (Exg1p) and chitinase (Cts1p), and (3) noncanonical proteins “classically” considered to be confined to the intracellular compartment because they lack the conventional secretory signal sequence (2,9–11). Nevertheless, the mechanism by which these nonconventional proteins are targeted to the cell surface remains enigmatic (11–13). This array of loosely associated proteins is commonly found at the cell surface but also, in a smaller ratio, in the cell-wall inner layers. These CWPs can be extracted using detergents and reducing agents (see Fig. 1). 2. Covalently linked to -1,3-glucan: a. Directly via an alkali-labile linkage (speculatively through a O-linked sidechain), such as PIR-CWPs (CWPs with internal repeats). The PIR-CWPs are highly O-mannosylated proteins with one or more internal repeat regions, a N-terminal signal peptide, a Kex2 proteolytic processing site, and a C-terminal sequence with four cysteine residues at highly conserved positions (1,14–17). They are normally located in the cell-wall inner layer (18). Other CWPs belong to this category are also present in the yeast cell walls (see the following and Fig. 2). This type of CWPs can be solubilized under mild alkali conditions or by enzymatic treatment with -1,3-glucanases but not with -1,6-glucanases (see Figs. 1–3). b. Indirectly by a -1,6-glucan moiety through their GPI remnant, such as GPICWPs. The GPI-CWPs are highly O-glycosylated proteins with an N-terminal signal peptide, a C-terminal GPI anchor addition signal, and serine- and threonine-rich regions (see Note 3) (1,7,17,19). These CWPs are predominantly placed in the cell-wall outer layer. This group of CWPs can be released either by enzymatic treatment with -1,3 or -1,6-glucanases or by using hydrofluoric acid (HF)-pyridine, which cleavages the phosphodiester bond in the GPI remnant (20) (GPI-CWP 1 in Figs. 1–3). 3. Covalently anchored to chitin by a -1,6-glucan moiety via their GPI remnant, such as some GPI-CWPs (21,22). This type of CWP-polysaccharide complex is largely found in the lateral walls or under stress conditions. These CWPs can be

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Fig. 2. Schematic representations of the different known types of covalently linked CWPs in S. cerevisiae and C. albicans, and the most commonly used methods to solubilize them from isolated cell walls. These diagrams are based on information from references (1,14,18-21,23). See Introduction for further details. PIR-CWPs, a small number of hybrid GPI-CWPs (e.g., Cwp1p (23); GPI-CWPs 2 and 3) at acidic pHs, and other CWPs (25,32) are directly attached to -1,3-glucan through an alkali-sensitive linkage (ASL). GPIr denotes GPI remnant, and ASL alkali-sensitive linkage.

extracted either by enzymatic treatment with chitinases or -1,6-glucanases or by using HF-pyridine (GPI-CWP 4 in Figs. 1–3).

However, it is unsurprising that other types of linkages, hitherto uncharacterized, among CWPs and structural wall components are also present in the yeast cell wall (see Fig. 1). Be that as it may, there is no doubt that the molecular model of the yeast cell wall is even more sophisticated than that outlined above. This is because certain CWPs can simultaneously be retained into the -1,3glucan/chitin skeleton in various ways. For instance, the Cwp1p, a S. cerevisiae GPI-CWP, is double-anchored to the -1,3-glucan framework both through an alkali-sensitive linkage and by its -1,6-glucan moiety, playing an important role in stress response (23). Hence, two additional GPI-CWP-polysaccharide complexes are defined (GPI-CWPs 2 and 3 in Figs. 2 and 3). Taking into account the special architecture and nature of the cell wall, the isolation and solubilization of CWPs from this complex cellular compartment is not therefore an evident and easy affair. Indeed, CWPs from yeasts

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Fig. 3. Venn diagram summarizing the most commonly used methods to extract the different known types of CWPs in S. cerevisiae and C. albicans. This chart is based on data from references (1,10,14,18–21,23). The GPI-CWP numbers refer to those indicated in Fig. 2.

and filamentous fungi are tricky enough to resolve by two-dimensional electrophoresis (2-DE) gels, because of their low abundance, low solubility, hydrophobicity, high heterogeneity, extensive glycosylation (especially, Oand/or N-mannosylation), and covalent attachment to the wall polysaccharide (-1,3-glucan/chitin) skeleton (11,24). These problems can, at least to some degree, be solved by sequential solubilization of CWPs on the basis of the type of attachments that they establish to other cell wall components (11,17,25). This procedure implies breakage of the covalent linkages between CWPs and wall polysaccharides (see Chapter 20). Intriguingly, cell wall fractionation is an appropriate paradigm system to: 1. Reduce the intricacy of the cell wall. 2. Enrich samples for CWPs and thus increase the detection of low-abundance species by removing the most abundant soluble gene products. 3. Enhance the solubility of large, low abundance, and/or hydrophobic CWPs. 4. Define (map and identify) the proteins that make up the cell wall (the cell wall proteome), and elaborate a comprehensive and integrated view of the complex CWP composition. The CWP resolution can be increased using the cell wall fractionation procedure described here, because the heterogeneous population of protein species present in the yeast and fungal cell envelope can be distributed over several 2-DE gels (11). 5. Characterize mechanisms of incorporation, assembly and retention of CWPs into the cell wall. 6. Elucidate the CWP interactions with cell wall polysaccharides.

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7. Study protein-protein interactions or regulatory networks exclusive to the cell wall. 8. Monitor abnormal protein expression localized to this external envelope. 9. Discover novel diagnostic/prognostic markers, antifungal targets and/or therapeutic candidates for human mycoses.

This chapter will integrate a typical procedure of cell wall fractionation to extract different CWP species from isolated cell walls of yeasts or filamentous fungi according to their interactions with other wall components. The resulting selectively enriched CWP fractions can then directly be (1) analyzed by 2DE (see Note 4) and mass spectrometry (MS) (11,17) or (2) digested with trypsin, followed by liquid chromatography (LC) in tandem with MS analyses (25) to circumvent some of the difficulties associated with in-gel digestion of the heavily glycosylated CWPs (12). The purity and quality of these enriched fractions of CWPs should be screened by using bona fide markers both of the cell wall and of intracellular compartments before carrying out any interpretation of the results. However, the unambiguous evidence for their cell wall location will only be established after (1) their in situ immunolocalization (i.e., immunoelectron microscopy or immunofluorescence studies) and/or (2) the use of tagged fusion proteins (e.g., c-myc-tag or green fluorescence protein (GFP) fusion proteins) based on the fusion of the protein in question to modified versions of extracellular enzymes that rely on a detectable phenotype.

2. Materials Growth media, solutions and buffers should be sterilized by autoclaving before use when working under sterile conditions. Their labile components should be sterilized separately using a 0.22-μm filter (Millipore, Bedford, MA) and then added to the other autoclaved ingredients. All solutions and buffers should be prepared with ultrapure water, as provided by Nanopure or Milli-Q 18 M/cm resistivity systems (Millipore), and precooled when the procedure is carried out at 4°C. 2.1. Cell Wall Isolation from Yeasts and Filamentous Fungi 1. Yeast-Peptone-D-glucose (YPD) plates: 1% (w/v) yeast extract (Difco Laboratories, Detroit, MI), 2% (w/v) peptone (Difco), 2% (w/v) d-glucose, 2% (w/v) agar (Difco). 2. YPD medium: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) d-glucose. 3. Bead mill homogenizer (model MSK; Braun Biotech International GmbH, Melsungen, Germany). 4. Liquid carbon dioxide (CO2 ).

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5. 0.40- to 0.60-mm chilled, acid-washed glass beads (Sartorius, Goettingen, Germany) (see Note 5). 6. PMSF stock solution: 0.1 M in isopropanol. Dissolve 174 mg of phenylmethylsulfonyl fluoride (PMSF; Fluka, Chelmsford, MA) in a final volume of 10 mL isopropanol, and store at –20°C (see Note 6). PMSF should be handled with caution because it is highly toxic. Weigh this hazardous compound in a fume hood, and wear gloves, goggles and a mask. 7. Lysis buffer: 10 mM Tris-HCl, pH 7.4, 1 mM PMSF. 8. YPD-chloramphenicol plates: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) d-glucose, 2% (w/v) agar (Difco), 10-μg/mL chloramphenicol. When the 1-L autoclaved YPD and agar solution cools to ∼65°C, add 1 mL of 10× chloramphenicol solution. 9. Chloramphenicol solution (10X): Dissolve 10 mg of chloramphenicol in a final volume of 1 mL ethanol, and sterilize using a 0.2-μm filter. 10. Wash solution A: 1 mM PMSF. 11. Wash solution B: 5% (w/v) NaCl, 1 mM PMSF. 12. Wash solution C: 2% (w/v) NaCl, 1 mM PMSF. 13. Wash solution D: 1% (w/v) NaCl, 1 mM PMSF.

2.2. Protein Solubilization from Isolated Yeast and Fungal Cell Walls 2.2.1. By Detergents and Reducing Agents 1. Wash buffer: 50 mM Tris-HCl, pH 8.0, 1 mM PMSF. 2. Extraction buffer: 50 mM Tris-HCl, pH 8.0, 0.1 M EDTA, 2% (w/v) SDS, 10 mM DTT (dithiothreitol; see Note 7).

2.2.2. Under Mild Alkali Conditions 1. 2. 3. 4.

Wash solution: 1 mM PMSF. Wash buffer: 0.1 M sodium acetate, pH 5.5, 1 mM PMSF. Extraction solution: 30 mM NaOH, 1 mM PMSF. Stop solution: acetic acid.

2.2.3. By -1,3-Glucanase Treatment 1. Wash solution: 1 mM PMSF. 2. Wash buffer: 50 mM Tris-HCl, pH 7.5, 1 mM PMSF. 3. Extraction buffer: 1,500 U Quantazyme ylgTM (Quantum Biotechnologies Inc, Montreal, Canada; see Note 8) per gram of wet weight of cell walls, in 2 mL of a solution containing 50 mM Tris-HCl, pH 7.5, 10 mM DTT, 1 mM PMSF (see Note 9). 4. Stop solution: 10% SDS.

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2.2.4. By Exochitinase Treatment 1. Wash solution: 1 mM PMSF. 2. Wash buffer: 50 mM sodium phosphate buffer, pH 6.3. 3. Extraction buffer: 0.3 U exochitinase (Sigma, St. Louis, MO) per gram of wet weight of cell walls, in 2 mL of a solution containing 50 mM sodium phosphate buffer, pH 6.3 (see Note 10). 4. Stop solution: 10% SDS.

2.3. Protein Precipitation. 1. 100% Trichloroacetic acid (TCA) solution: Dissolve 100 g of TCA in sufficient water to yield a final volume of 100 mL (see Note 11). TCA should be handled with caution, because it is extremely caustic. Protect eyes and avoid contact with skin when working with TCA solutions. 2. Acetone precooled to –20°C. 3. Neutralizing solution: 0.1 N NaOH.

3. Methods The protocols described below outline a typical procedure of cell wall fractionation to isolate and solubilize different CWP species from yeasts and filamentous fungi on the basis of the type of linkages that they establish with other wall components. This method involves (1) cell homogenization by physical disruption techniques, (2) isolation of cell walls by differential centrifugation, (3) sequential solubilization of CWPs from isolated cell walls using different chemical agents (detergents, reducing agents, and alkalis) and enzymes (glucanases and chitinases), and (4) precipitation of the resulting selectively enriched CWP fractions under suitable conditions for subsequent proteomic analyses. A flowchart of the strategy presented here is shown in Fig. 4. This is based on earlier methods described by Kapteyn et al. (20,21) and Mrsa et al. (26), and on recent modifications reported by Pitarch et al. (11) that have proved effective at properly extracting CWPs from S. cerevisiae and C. albicans. Nevertheless, adaptation of growth and extraction conditions may be required for other species of yeasts and filamentous fungi. 3.1. Cell Wall Isolation from Yeasts and Filamentous Fungi (see Note 12) Although cell disintegration can be accomplished using a wide variety of techniques, mechanical breakage of cells using glass beads is certainly one of the most quick and reliable procedures to disrupt the cell walls and plasma membranes of yeasts and filamentous fungi (see Note 13). After cell disruption,

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Fig. 4. Flowchart of a typical procedure of cell wall fractionation to isolate and solubilize different CWP species from yeasts and filamentous fungi according to the type of attachments that they establish to other cell wall components.

cell walls can be separated from other cytosolic and membranous components by differential centrifugation of the cell homogenate at relatively low speed (see Note 14).

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1. Grow yeast or fungal cells on an YPD plate (stock maintenance medium) at 30°C for 2 days. Use a single colony to inoculate 50 mL of YPD (or selective) medium in a 250-mL flask (see Note 15), and grow overnight at 30ºC with vigorous rotary shaking (200 rpm). 2. Use this 50-mL preculture to inoculate four 2-L flasks containing 500 mL fresh YPD medium each, and grow at 30°C in a shaking incubator (200 rpm) until the culture reaches early-log phase growth (OD600nm = 0.5–1; see Note 16). 3. Harvest the cells by centrifugation at 4,500g for 5 min (see Note 17) and discard the supernatant. 4. Resuspend the cell pellet in 200 mL ice-cold water, and centrifuge 5 min at 4,500g. Decant the supernatant. 5. Resuspend the cell pellet in 200 mL ice-cold lysis buffer, and centrifuge 5 min at 4,500g. Discard the supernatant. 6. Resuspend the cells in 3 volumes of ice-cold lysis buffer, and add 3–4 volumes of 0.5-mm acid-washed glass beads (see Note 18). Transfer the cell suspension to an appropriate sized shaking flask (see Note 19). 7. Grind the suspension at maximum speed for 30–60 s using a bead mill homogenizer and cooling with liquid CO2 (for example, in a CO2 -refrigerated MSK homogenizer), and then place the shaking flask on ice for 1–2 min (see Note 20). Repeat this step until complete cell breakage. Monitor the degree of cell breakage with a phase-contrast microscope and by plating on YPD-chloramphenicol plates (see Note 21). 8. Enable the glass beads to settle out and collect the supernatant carefully (see Note 22). Wash the glass beads with ice-cold lysis buffer and collect the washings until they are clear (see Note 23). Pool the supernatant and all the washings. 9. Centrifuge the pooled supernatant and washings (cell homogenate) at 1,000g for 10 min at 4°C. Discard the supernatant carefully (see Note 24). 10. Resuspend the isolated cell walls in 200 mL ice-cold wash solution A. Centrifuge 10 min at 1,000g and at 4°C. Carefully decant the supernatant. Repeat this step two more times. 11. Resuspend the cell walls in 200 mL ice-cold wash solution B. Centrifuge 10 min at 1,000g and at 4°C. Carefully decant the supernatant. Repeat this step four more times (see Note 25). 12. Resuspend the walls in 200 mL ice-cold wash solution C. Centrifuge 10 min at 1,000g and at 4°C. Carefully remove the supernatant. Repeat this step four more times. 13. Resuspend the pellet in 200 mL ice-cold wash solution D. Centrifuge 10 min at 1,000g and at 4°C. Carefully decant the supernatant. Repeat this step four more times (see Note 26). 14. Resuspend the walls in 200 mL ice-cold wash solution A. Centrifuge 10 min at 1,000g and at 4°C. Carefully discard the supernatant. Repeat this step two more times using preweighed centrifuge bottles. 15. Weigh the wet wall pellet (see Note 27).

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3.2. Protein Solubilization from Isolated Yeast and Fungal Cell Walls CWPs can then be solubilized sequentially from isolated cell walls by a wide variety of reagents (e.g., detergents, reducing agents, alkalis, and hydrolytic enzymes, among others) in connection with their attachments to other wall components (see Fig. 4). All procedures are performed at 4°C with prechilled solutions, reagents and apparatus (see Note 28) unless otherwise indicated. 3.2.1. By Detergents and Reducing Agents Detergents, such as sodium dodecyl sulfate (SDS) or n-octylglucoside, can be used to extract CWPs that are associated noncovalently with other wall components. The use of reducing agents, such as dithiothreitol (DTT) or mercaptoethanol (ME), (1) enables the solubilization of CWPs that are loosely associated, either by disulfide bridges or through non-covalent bonds, with other covalently linked CWPs, (2) results in an increase in the cell wall porosity, and (3) facilitates the subsequent action of wall degrading enzymes (see Figs. 1 and 4). 1. Resuspend the purified cell walls (prepared as described in Subheading 3.1) in 200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g and carefully decant the supernatant (see Note 24). 2. Add 5 mL of extraction buffer for each wet gram of cell walls and resuspend. Boil the cell wall suspension at 100°C for 10 min, and centrifuge 10 min at 1,000g. 3. Collect the supernatant and store the pellet. 4. Precipitate the supernatant (containing SDS/DTT-extractable CWPs) and store at –80°C (see Note 29 and Fig. 4). 5. Repeat step 2 with the stored pellet and carefully discard the supernatant (see Note 30). 6. Weigh the wet wall pellet (containing SDS/DTT-resistant cell walls) and divide it equally between two tubes (see Fig. 4 and Notes 27 and 31). Store them for further extractions (see Subheadings 3.2.2, 3.2.3 and 3.2.4).

3.2.2. Under Mild Alkali Conditions Treatment of SDS/DTT-resistant cell walls under mild alkali conditions (using 30 mM NaOH) results in the extraction of CWPs linked to -1,3-glucan through an alkali-sensitive linkage (of unknown nature) by the -elimination process (1, 14–17). PIR-CWPs, some GPI-CWPs, and other CWPs belong this group (see Figs. 1–4). 1. Resuspend one tube of the purified SDS/DTT-resistant cell walls (prepared as described in Subheading 3.2.1) in 200 mL ice-cold wash solution. Centrifuge 10 min at 1,000g and carefully decant the supernatant (see Note 24). Repeat this step two more times. 2. Resuspend the walls in 200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g and carefully discard the supernatant. Repeat this step five to seven more times.

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3. Add 4 mL of ice-cold extraction solution for each wet gram of cell walls and resuspend. Incubate the cell wall suspension at 4°C for 17 h with gentle shaking (see Note 32). 4. Stop the chemical reaction by adding neutralizing amounts of acetic acid (see Note 33). 5. Centrifuge 10 min at 1,000g and collect the supernatant (containing alkalisensitive CWPs). Precipitate or dialyze the clear supernatant (see Note 33) and store at –80°C.

3.2.3. By -1,3-Glucanase Treatment -1,3-glucanases (commercially available) can be used to extract CWPs covalently anchored to -1,3-glucan. This group of CWPs released from SDS/DTT-resistant cell walls by these wall hydrolytic enzymes contains (1) GPI-CWPs, which are indirectly attached to -1,3-glucan, via -1,6-glucan, through a phosphodiester bridge and can alternatively be solubilized either by 1,6-glucanases (see Note 34) or by using HF-pyridine, (2) alkali-sensitive CWPs (including PIR-CWPs and some GPI-CWPs, among others), which are anchored to -1,3-glucan through uncharacterized linkages (perhaps by a O-linked sidechain) and can also be released under mild alkali conditions (using 30 mM NaOH; see Subheading 3.2.2), and (3) other CWPs linked to -1,3-glucan through other types of hitherto unidentified bridges (see Figs. 1–4). 1. Resuspend the other tube of the purified SDS/DTT-resistant cell walls (prepared as described in Subheading 3.2.1) in 200 mL ice-cold wash solution. Centrifuge 10 min at 1,000g and carefully decant the supernatant (see Note 24). Repeat this step two more times. 2. Resuspend the walls in 200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g and carefully remove the supernatant. Repeat this step five to seven more times. 3. Add 2 mL of extraction buffer for each wet gram of cell walls and resuspend (see Note 34). Incubate the cell wall suspension at 37°C for 17 h with gentle shaking. 4. Stop the enzymatic reaction by adding the stop solution at a final concentration of 0.4% (w/v) and heating at 100°C for 3–5 min. 5. Centrifuge 10 min at 1,000g. 6. Collect the supernatant (containing -1,3-glucanase-extractable CWPs) and store the pellet (containing SDS/DTT and -1,3-glucanase-resistant cell walls) for further extractions (see Subheading 3.2.4). 7. Precipitate the clear supernatant and store at –80°C.

3.2.4. By Exochitinase Treatment Enzymatic treatment of the SDS/DTT- and -1,3-glucanase-resistant cell walls with exochitinases leads to the solubilization of CWPs covalently anchored to chitin. These comprise (1) a small subgroup of GPI-CWPs, which

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are indirectly linked to chitin, via -1,6-glucan, through their GPI remnant and can also be extracted either by -1,6-glucanases or by using HF-pyridine, and (2) other CWPs attached to chitin through other types of hitherto uncharacterized linkages (see Figs. 1–4). 1. Resuspend the SDS/DTT and -1,3-glucanase-resistant cell walls (prepared as described in Subheading 3.2.3) in 200 mL ice-cold wash solution. Centrifuge 10 min at 1,000g and carefully decant the supernatant (see Note 24). Repeat this step two more times. 2. Resuspend the walls in 200 mL ice-cold wash buffer. Centrifuge 10 min at 1,000g and carefully discard the supernatant. Repeat this step five to seven more times. 3. Add 2 mL of extraction buffer for each wet gram of cell walls and resuspend. Incubate the cell wall suspension at 37°C for 17 h with gentle shaking. 4. Stop the enzymatic reaction by adding the stop solution at a final concentration of 0.4% (w/v) and heating for 3–5 min at 100°C. 5. Centrifuge 10 min at 1,000g and collect the supernatant (containing -1,3-glucanase-resistant and exochitinase extractable CWPs). 6. Precipitate the clear supernatant and store at –80°C.

3.3. Protein Precipitation. The different selectively enriched CWP fractions obtained in the Subheading 3.2 should be concentrated and desalted before carrying out further proteomic analyses. These can be (1) dialyzed against a volatile buffer and dried, or (2) precipitated with TCA as described below (see Note 35), among other methods. 1. Add 1/9th the total volume of the protein sample of an ice-cold 100% TCA solution to a final concentration of 10%. 2. Mix thoroughly and incubate on ice for 30 min. 3. Centrifuge the suspension at 10,000g for 15 min, and discard the supernatant (see Note 36). 4. Wash protein pellet twice with cold acetone to remove residual TCA. 5. Air dry for 30 min. 6. Add neutralizing amounts of 0.1N NaOH (see Note 37) and store at –80°C.

4. Notes 1. Supporting this enterprise, -1,3-glucan is targeted by a new antifungal drug class in recent clinical use, i.e., echinocandins (including caspofungin and micafungin), which blocks the biosynthesis of this cell wall polysaccharide (4–6). 2. It must be borne in mind that unlike S. cerevisiae, many filamentous fungi contain further -glucans and a high chitin content in their cell walls (1). 3. The GPI-proteins are translocated into the endoplasmic reticulum (ER), where (1) the N-terminal signal peptide (secretion signal necessary to enter the classical

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5. 6.

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ER-Golgi secretory pathway) is cleaved, (2) the C-terminal GPI anchor addition signal is replaced with a GPI anchor, and (3) O- and/or N-linked core glycosylation takes place. Remarkably, further glycosylation also then occurs in the Golgi apparatus. These GPI-anchored proteins are directed through the secretory pathway to the outer side of the plasma membrane, where they are attached through their C-terminal GPI anchors (see Fig. 5). Intriguingly, some of them are released from the plasma membrane by cleavage of their GPI anchors, resulting in GPI anchor remnants (a truncated, lipidless GPI anchors) (1,19,27,28). These proteins are then covalently incorporated into the cell wall, by the attachment of their GPI remnants to -1,6-glucan (19). It is convenient to use gradient gels that enable protein separation up to at least 500 kDa, because the extensive glycosylation (especially, O- and/or Nmannosylation) of a large proportion of CWPs results in extremely high apparent molecular masses on SDS-PAGE or 2-DE gels (7). The size of the glass beads is crucial to achieve an efficient cell disruption. Optimal bead size for spores is 0.1 mm and for yeast and mycelia 0.5 mm. The PMSF can also be solubilized in ethanol, methanol and 1,2-propanediol. It is unstable in aqueous solution. PMSF is added to reduce possible proteolytic processes. It inhibits serine proteases (e.g., trypsin, chymotrypsin, and thrombin) and thiolproteases (e.g., papain). 40 mM -mercaptoethanol may be substituted for 10 mM DTT. mercaptoethanol or DTT should be added just before use. Quantazyme ylgTM is a recombinant yeast -1,3-glucanase purified from E. coli that is completely free of protease, endo- and exonuclease activity. It is highly stable in solution for months at 4°C, maintaining its whole activity. The use of reducing agents facilitates the ability of Quantazyme ylgTM to degrade the cell wall -1,3-glucan. 40 mM -mercaptoethanol or 10 mM cysteine may be substituted for 10 mM DTT. Exochitinase is a cell wall lytic enzyme isolated from Serratia marcescens that catalyzes the progressive degradation of chitin, starting at its nonreducing end. This preparation contains phosphate buffer salts and shows an optimum pH of 6.0. Because TCA is very hygroscopic, the whole content of a newly opened TCA bottle should be used to prepare the TCA stock solution. Perform all procedures from this subheading until isolated cell walls are obtained under sterile conditions. Use sterile centrifuge bottles. Mechanical cell breakage using a bead mill homogenizer is considered the technique of choice for disrupting cells with cell walls, especially spores, yeasts and fungi, but it also works successfully with algae, bacteria, plant, and animal tissue culture in suspension. Cell disruption takes place by the crushing action of the glass beads, which are vigorously agitated by shaking or stirring, after crashing with the cells. The Braun MSK cell homogenizer combines two types of motions in the shaking flask: rotation and tumbling. For small-scale preparations, a Fast-Prep cell breaker (Q-Biogene, Carlsbad, CA) can be used in lieu

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Fig. 5. Putative model of the incorporation of GPI-anchored proteins into the cell wall. This is based on information from references (19,27,28). Their GPI anchor is processed at the plasma membrane leading to a GPI remnant (a truncated, lipidless GPI anchor). This is cross-linked to -1,6-glucan when these GPI proteins are covalently incorporated into the cell wall. See Note 3 for further details.

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of the Braun MSK cell homogenizer. Cell breakage of filamentous fungi can alternatively be achieved by (1) grinding freeze-dried mycelium in a mortar and pestle, (2) grinding mycelium in liquid nitrogen, or (3) homogenizing thick cell pastes. When handling liquid nitrogen, use insulated gloves, avoid any potential contact because of the risk of frostbite, and never utilize glass containers because they may break. Differential centrifugation in sucrose density gradients may alternatively be used to isolate the cell wall fraction. Liquid cultures should be grown in a flask that is at least 4–5 times larger than the culture volume. It is important that the yeast culture is in early-log phase growth (∼ OD600nm = 0.5–1) because it is easier to disrupt their cell walls with the bead mill homogenizer than those close to or in stationary phase growth. It must be borne in mind that the composition of the cell wall changes with the growth stage and culture conditions (growth temperature, external pH, oxygen levels, and composition of the growth medium (1,29)), among others. Hence, it may be necessary to adjust the cell density according to the specific objectives of the experiment. Overall, yeasts are successfully harvested by centrifugation, whereas vacuumassisted filtration, rather than centrifugation, is often preferred for harvesting filamentous fungi. The wet weight (in grams) of the cell pellet is nearly equal to the packed cell volume (in milliliters). Add about 3 mL of ice-cold lysis buffer for each wet gram of cell pellet and resuspend. The shaking flasks can be made of glass or stainless steel. The latter are better at transferring heat. It is essential to exclude all air from the shaking flask before the cell breakage to avoid foaming and denaturation of proteins. It is extremely important that the temperature of the cell suspension remains at 4°C during the cell disruption to prevent heat inactivation and denaturation of proteins. Use liquid CO2 to cool the protein sample during cell breakage. This procedure is carried out until complete cell breakage, which will vary with yeast/fungal strain and growth stage. The degree of cell breakage should be examined: a. Before proceeding: by observing cell lysis under a phase-contrast microscope. b. After proceeding: by plating an aliquot of the cell suspension after and before cell disruption on YPD-chloramphenicol plates and growing at 30°C. The ratio of CFUs after to before cell breakage is then calculated to estimate the efficiency of cell disruption and, therefore, potential intracellular contamination in the subsequent steps. Failure of cells to grow on YPD-chloramphenicol plates should be evidenced.

22. The supernatant (cell homogenate) and following washings (residual cell homogenate) can be collected (1) by decanting after allowing the glass beads to settle out by gravity, or (2) by straining through a perforated tube or “strainer”

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Pitarch et al. (see Fig. 6). To perform the last method, transfer the cell homogenate containing the glass beads to a plastic tube and make holes of a diameter less than 0.4–0.6 mm in its bottom with a flamed needle to obtain a “strainer.” After straining the cell homogenate off, collect it and wash the glass beads with ice-cold lysis buffer. Collect the washing, and wash again until the collected washings are clear. Pool the cell homogenate and all the washings. The washing step is critical

Fig. 6. Proposed procedure for removing the glass beads from the cell homogenate. In the method described here, the cell homogenate is strained through a perforated tube with holes of a diameter less than 0.4–0.6 mm in its bottom, whereas the glass beads (with a diameter of 0.4–0.6 mm) are retained in the perforated tube or “strainer” (see Note 22). Successive washings with ice-cold lysis buffer should then be performed to increase the recovery of cell homogenate, and thus of cell wall material.

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to avoid any loss of material while removing the glass beads from the cell homogenate. To reuse the glass beads, rinse them by soaking in concentrated nitric acid for 1 h, and then flush through with water. Dry them in a baking oven, cool, and store at 4°C (see Fig. 6). It is important to decant the supernatant carefully, because the cell-wall pellet is less compact than the preceding cell pellets. In general, care should be taken during the following steps of this procedure (especially during decanting of the washings) to prevent any loss of cell walls, and therefore of CWPs. Be that as it may, this potential loss of protein material does not interestingly result in a preferential deficiency of certain CWP species or families. The purpose of extensively washing the isolated cell walls with solutions of decreasing concentrations of NaCl is to remove potential extracellular, membranous and/or cytosolic protein contaminants that can be adhered to them through nonspecific ionic interactions (see Fig. 7). The number of washing steps will therefore rely on the objectives of the experiment. A further washing step with 0.1 M Na2 CO3 overnight at 4°C under gentle shaking may reduce potential cytoplasmic contamination that is found into microsomes, because this treatment allows them to be opened and washed (30,31). The wet weight (in grams) of cell walls in the pellet can be calculated by taking the weight of the centrifuge bottle (which has been previously weighed) away from the total weight (i.e., the weight of wall pellet plus centrifuge bottle). Centrifugation must be performed in refrigerated centrifuges at 4°C, with prechilled rotors, to avoid undesirable proteolytic activity. This fraction is not a pure preparation of cell wall, but rather is enriched in CWPs (loosely associated with other wall components) and may potentially contain a small amount of membrane proteins. This extra step is important to remove any remaining SDS/DTT-extractable CWPs and potential membranous components from the isolated cell walls, which will be used in subsequent CWP extraction steps (see Fig. 4). This pellet, containing SDS/DTT-resistant cell walls, is divided equally between two tubes to independently extract alkali-sensitive CWPs and -1,3-glucanaseextractable CWPs in the following steps (see Fig. 4). It is convenient to place the cell wall suspension into a container with ice in a cool room at 4°C (see Fig. 4). The chemical reaction must be stopped by acid neutralization. This can be performed: a. by adding neutralizing amounts of acetic acid to the wall suspension. The clear supernatant containing alkali-sensitive CWPs should then be (i) precipitated by adding nine volumes of ice-cold methanol (18) or (ii) dialyzed against water or 20 mM bis-Tris, pH 6.0 (32). b. by subsequent protein precipitation of the clear supernatant containing alkalisensitive CWPs with TCA (see Subheading 3.3).

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Fig. 7. Basic principle of the procedure used for washing the isolated cell walls of yeasts and filamentous fungi. Potential extracellular, membranous and/or cytosolic protein contaminants that may nonspecifically associate with the isolated cell walls through ionic interactions can be removed under relatively more stringent conditions. Extensive washings of the isolated cell walls with high ionic strength (e.g., 5% NaCl) can disrupt the nonspecific ionic interactions between the isolated cell walls and these putative contaminants.

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34. Endo--1,6-glucanase isolated from Trichoderma harzianum (33) can be used to release GPI-CWPs (0.8 U/g wet weight of cell walls in 100 mM sodium acetate pH 5.5) (18,20). Subsequently, -1,6-glucanase-digested cell walls can be treated with Quantazyme or ice-cold 30 mM NaOH to extract (1) the 1,6-glucanase-resistant PIR-CWPs, (2) the -1,6-glucanase-resistant GPI-CWPs (GPI-CWPs linked to the -1,3-glucan through a alkali-sensitive linkage), and (3) hybrid GPI-CWPs, such as Cwp1p (see Fig. 2) (23). 35. The protein concentration should be higher than 100 μg/mL before precipitating with TCA. If the amount of protein precipitated is less than 1 nmole, the protein sample should be concentrated and desalted by other ways (e.g., by ultrafiltration or forced dialysis), because the pellet is imperceptible. 36. The supernatant should be stored in case the protein did not precipitate. 37. Protein preparation can then be resuspended in a small volume of buffer suitable for the subsequent analytical procedures (e.g. sample buffer for SDS-PAGE or 2-DE).

Acknowledgments We thank the Merck, Sharp & Dohme (MSD) Special Chair in Genomics and Proteomics, European Community (STREP LSHB-CT-2004-511952), Comunidad de Madrid (S-SAL-0246/2006) and Comisión Interministerial de Ciencia y Tecnología (BIO-2003-00030 and BIO-2006-01989) for financial support of our laboratory. References 1. Klis, F. M., Boorsma, A. and de Groot, P. W. (2006) Cell wall construction in Saccharomyces cerevisiae. Yeast, 23, 185–202. 2. Chaffin, W. L., Lopez-Ribot, J. L., Casanova, M., Gozalbo, D. and Martinez, J. P. (1998) Cell wall and secreted proteins of Candida albicans: identification, function, and expression. Microbiol Mol.Biol Rev. 62, 130–80. 3. Bernard, M. and Latge, J. P. (2001) Aspergillus fumigatus cell wall: composition and biosynthesis. Med.Mycol. 39, 9–17. 4. Odds, F. C., Brown, A. J., and Gow, N. A. (2003) Antifungal agents: mechanisms of action. Trends Microbiol. 11, 272–79. 5. Morrison, V. A. (2006) Echinocandin antifungals: review and update. Expert.Rev.Anti.Infect.Ther. 4, 325–42. 6. Deresinski, S. C. and Stevens, D. A. (2003) Caspofungin. Clin.Infect.Dis. 36, 1445–57. 7. de Groot, P., Ram, A. F., and Klis, F. M. (2005) Features and functions of covalently linked prtoeins in fungal cell walls. Fungal.Genet.Biol. 42, 657–75. 8. Ruiz-Herrera, J., Elorza, M. V., Valentin, E and Sentandreu, R. (2006) Molecular organization of the cell wall of Candida albicans and its relation to pathogenicity. FEMS Yeast Res. 6, 14–29.

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9. Klis, F. M., de Groot, P. and Hellingwerf, K. (2001) Molecular organization of the cell wall of Candida albicans. Med.Mycol. 39, 1–8. 10. Cappellaro, C., Mrsa, V. and Tanner, W. (1998) New potential cell wall glucanases of Saccharomyces cerevisiae and their involvement in mating. J.Bacteriol. 180, 5030–37. 11. Pitarch, A., Sanchez, M., Nombela, C. and Gil, C. (2002) Sequential fractionation and two-dimensional gel analysis unravels the complexity of the dimorphic fungus Candida albicans cell wall proteome. Mol.Cell Proteomics 1, 967–82. 12. Pitarch, A., Nombela, C., and Gil, C. (2006) Candida albicans biology and pathogenicity: Insights from proteomics, in Microbial Proteomics: Functional Biology of Whole Organisms (Humphery-Smith,I. and Hecker,M., eds), Wiley-Vch, Hoboken, NJ, pp. 285–330. 13. Nombela, C., Gil, C. and Chaffin, W. L. (2005) Non-conventional protein secretion in yeast. Trends Microbiol. 14, 15–21. 14. Mrsa, V. and Tanner, W. (1999) Role of NaOH-extractable cell wall proteins Ccw5p, Ccw6p, Ccw7p and Ccw8p (members of the Pir protein family) in stability of the Saccharomyces cerevisiae cell wall. Yeast 15, 813–20. 15. Kapteyn, J. C., Van Egmond, P., Sievi, E., Van Den, E. H., Makarow, M. and Klis, F. M. (1999) The contribution of the O-glycosylated protein Pir2p/Hsp150 to the construction of the yeast cell wall in wild-type cells and beta-1,6-glucandeficient mutants. Mol.Microbiol. 31, 1835–44. 16. Kandasamy, R., Vediyappan, G. and Chaffin, W. L. (2000) Evidence for the presence of Pir-like proteins in Candida albicans. FEMS Microbiol Lett. 186, 239–43. 17. Weig, M., Jansch, L., Gross, U., De Koster, C. G., Klis, F. M., and de Groot, P. W. (2004) Systematic identification in silico of covalently bound cell wall proteins and analysis of protein-polysaccharide linkages of the human pathogen Candida glabrata. Microbiology 150, 3129–44. 18. Kapteyn, J. C., Hoyer, L. L., Hecht, J. E., et al. (2000) The cell wall architecture of Candida albicans wild-type cells and cell wall-defective mutants. Mol.Microbiol. 35, 601–11. 19. Kollar, R., Reinhold, B. B., Petrakova, E., et al. (1997) Architecture of the yeast cell wall. -1,6-glucan interconnects mannoprotein, -1,3-glucan, and chitin. J.Biol Chem. 272, 17762–75. 20. Kapteyn, J. C., Montijn, R. C., Vink, E., et al. (1996) Retention of Saccharomyces cerevisiae cell wall proteins through a phosphodiester-linked -1,3-/-1,6-glucan heteropolymer. Glycobiology 6, 337–45. 21. Kapteyn, J. C., Ram, A. F., Groos, E. M., et al. (1997) Altered extent of cross-linking of -1,6-glucosylated mannoproteins to chitin in Saccharomyces cerevisiae mutants with reduced cell wall -1,3-glucan content. J.Bacteriol. 179, 6279–84. 22. Sestak, S., Hagen, I., Tanner, W. and Strahl, S. (2004) Scw10p, a cell-wall glucanase/transglucosidase important for cell-wall stability in Sacccharomyces cerevisiae. Microbiology, 150, 3197–3208.

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23. Kapteyn, J. C., ter Riet, B., Vink, E., et al. (2001) Low external pH induces HOG1dependent changes in the organization of the Saccharomyces cerevisiae cell wall. Mol.Microbiol. 39, 469–479. 24. Pitarch, A., Sanchez, M., Nombela, C. and Gil, C. (2003): Analysis of the Candida albicans proteome. I. Strategies and applications. J.Chromatogr.B Analyt.Technol.Biomed.Life Sci. 787, 101–128. 25. de Groot, P. W., de Boer, A. D., Cunningham, J., et al. (2004) Proteomic analysis of Candida albicans cell walls reveals covalently bound carbohydrate-active enzymes and adhesins. Eukaryot.Cell 3, 955–65. 26. Mrsa, V., Seidl, T., Gentzsch, M. and Tanner, W. (1997) Specific labelling of cell wall proteins by biotinylation. Identification of four covalently linked O-mannosylated proteins of Saccharomyces cerevisiae. Yeast 13, 1145–54. 27. Lipke, P. N. and Kurjan, J. (1992) Sexual agglutination in budding yeasts. Structure, function, and regulation of adhesion glycoproteins. Microbiol. Rev. 56, 180–94. 28. Roh, D. H., Bowers, B., Riezman, H. and Cabib, E. (2002) Rho1p mutations specific for regulation of -1,3-glucan synthesis and the order of assembly of the yeast cell wall. Mol.Microbiol. 44, 1167–83. 29. Aguilar-Uscanga, B. and Francois, J. M. (2003) A study of the yeast cell wall composition and structure in response to growth conditions and mode of cultivation. Lett.Appl.Microbiol. 37, 268–74. 30. Fujiki, Y., Hubbard, A. L., Fowler, S. and Lazarow, P. B. (1982) Isolation of intracellular membranes by means of sodium carbonate treatment: application to endoplasmic reticulum. J.Cell Biol. 93, 97–102. 31. Lopez-Villar, E., Monteoliva, L., Larsen, M. R., et al. (2006) Genetic and proteomic evidences support the localization of yeast enolase in the cell surface. Proteomics 6, S107–S118. 32. Yin, Q. Y., de Groot, P., Dekker, H. L., de Jong, L., Klis, F. M. and de Koster, C. G. (2005) Comprehensive proteomic analysis of Saccharomyces cerevisiae cell walls: Identification of proteins covalently attached via glycosylphosphatidylinositol remnants or mild alkali-sensitive linkages. J.Biol Chem. 280, 20894–901. 33. De La Cruz, J., Pintor-Toro, J. A., Benitez, T. and Llobell, A. (1995) Purification and characterization of an endo--1,6-glucanase from Trichoderma harzianum that is related to its mycoparasitism. J.Bacteriol. 177, 1864–71.

20 Collection of Proteins Secreted from Yeast Protoplasts in Active Cell Wall Regeneration Aida Pitarch, César Nombela, and Concha Gil

Summary The yeast cell wall is a dynamic and complex matrix of polysaccharides (glucans, mannans, and chitin), proteins and minor amounts of lipids that isolate the yeast from the extracellular medium, protecting it against osmotic and physical injuries. Removal of this essential structure for cell integrity and viability by controlled enzymatic digestion in an iso-osmotic medium brings about protoplast formation. When yeast protoplasts are incubated in an osmotically stabilized liquid nutrient medium, cell wall precursors are secreted into the culture medium to de novo synthesize the cell wall. During the early stages of the regeneration process of protoplast walls, many wall protein precursors (presumably structural proteins along with remodeling and cross-linking enzymes) are shed into the extracellular medium but not covalently incorporated into the nascent cell wall, intriguingly enabling their easy isolation and solubilization. We have developed a method to collect proteins secreted from yeast protoplasts in active cell wall regeneration under conditions that are suitable for subsequent proteomic analyses. This procedure circumvents some of the troubles intrinsically related to other extraction protocols of cell wall proteins, such as chemical or enzymatic modifications, and poor quality in protein resolution and identification because of linkages to glucan/chitin residues. It further offers a valuable model system to understand how the de novo cell wall biosynthesis occurs in the yeast cell or how the yeast cell wall participates in morphogenesis.

Key Words: Candida albicans; cell wall; protoplasts; regeneration; Saccharomyces cerevisiae; yeast.

1. Introduction Yeasts are unicellular eukaryotic organisms that, unlike mammalian cells, are surrounded by an elastic and highly dynamic structure, namely the cell wall. From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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This external envelope, basically composed of -1,3 and -1,6-glucans, chitin, mannoproteins, and proteins (1,2), is crucial for preserving the morphology and osmotic integrity of the yeast cell and, therefore, essential for cell viability (see chapter on Cell Wall Fractionation for Yeast and Fungal Proteomics for further details). The yeast cell wall can be completely eliminated by controlled enzymatic digestion in an iso-osmotic medium, resulting in the formation of protoplasts (see Fig. 1). Yeast protoplasts are uniquely enveloped by the plasma membrane, which displays typical invaginations and dictates their distinctive spherical shape on the basis of physical laws. When yeast protoplasts are cultivated in osmotically stabilized nutrient media, these are intriguingly able to synthesize a new cell wall and revert to normal cells, which are capable of proliferating and inducing proper morphogenesis (see Note 1) (3–6). Both processes, cell wall regeneration and protoplast reversion, are intrinsically associated with yeast cell survival (see Note 2) (5). Chitin is the first cell wall component to be deposited on the surface of regenerating protoplasts (7–9). The microfibrils of nascent chitin, distributed irregularly over the plasma membrane at the early stages of reversion, undergo a gradual rise in density during the subsequent stages of cell wall regeneration, leading to the formation of a regular fibrillar mesh. This de novo chitin skeleton, located around the plasma membrane of reverting protoplasts, is soon overlaid with -1,3-glucan. Remarkably, although cell wall proteins (i.e., mannoproteins and proteins) begin to be synthesized early in the course of protoplast reversion to normal cells, their covalent incorporation into this de novo glucan-chitin framework is delayed until an adequate amount of -1,3-glucan molecules is assembled on the nascent polysaccharide lattice. In fact, cell wall proteins are the last components to be effectively bound to the regenerating wall. For this reason, during the initial stages of the reversion process, cell wall proteins (biologically intended for being anchored to the wall polysaccharide network) are not covalently retained into the nascent wall of regenerating protoplasts and, consequently, are shed into the extracellular medium. A schematic representation of the dynamics of protoplast formation and de novo wall construction in reverting protoplasts is shown in Fig. 1. The use of regenerating protoplasts and, in particular the collection of proteins secreted from yeast protoplasts in active cell wall regeneration (i.e., during the first stages of the regeneration process of protoplast walls) into the culture medium, are a good and simple model system to: 1. Easily isolate and solubilize proteins of the complex yeast cell wall, because these are freely released from actively reverting protoplasts into the culture medium. 2. Reduce the complexity of this subcellular compartment (see Chapter 19 for details on intricate structure and molecular organization of the yeast cell wall) and

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Fig. 1. Dynamics of protoplast formation, cell wall regeneration and protoplast reversion to normal yeast cells. Elimination of the yeast cell wall by controlled enzymatic digestion in an iso-osmotic medium leads to the protoplast formation (see Note 14). Yeast protoplasts have typical intrinsic features, such as (1) spherical shape, (2) invaginations of the plasma membrane in many areas, and (3) ability to synthesize new cell walls and revert to normal growing cells in iso-osmotic regenerating conditions (see Note 1). Regenerating protoplasts in an osmotically stabilized nutrient medium

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facilitate further related analytical procedures, exploiting the fact that the yeast cell is deprived of its cell wall and stimulated to resynthesize it step by step. 3. Bypass some of the problems inherently associated with chemical and/or enzymatic treatments to solubilize cell wall proteins (see Chapter 19), such as: a. Potential protein modifications. These are avoided with the method described here because cell wall proteins are collected from the growth medium and directly analyzed in ensuing proteomic studies without the use of chemical agents or enzymes that may in some measure modify them. b. The extraction of proteins bearing glucan and/or chitin side-chain residues, which hinder their subsequent resolution by two-dimensional gel electrophoresis (2-DE) and/or identification by mass spectrometry (MS) (10, 11). This is circumvented using the present approach, because this does not break the covalent linkages between proteins and structural wall polysaccharides (glucan/chitin). 4. Characterize cell wall protein precursors before their incorporation into the cell wall (i.e., those gene expression products involved in the de novo wall biosynthesis). These mainly include structural proteins, as well as remodelling and cross-linking enzymes (10,12–14)). 5. Study the de novo generation and, therefore, all steps of the biosynthesis of the cell wall of yeasts, because this stratagem yields information not only about the composition of cell wall precursors but also about their interactions (at the different stages of protoplast reversion to normal cells) (7–9,12). 6. Elucidate the role of the yeast wall in cell morphogenesis, given that cell wall regeneration by reverting protoplasts is a de novo process of morphogenesis (3–5,15).

Fig. 1. (Continued) first form a fibrillar chitin network, whereupon -1,3-glucan molecules are promptly deposited. Mannoproteins and proteins are the last components to be assembled on the nascent fibrillar mesh, because their covalent incorporation into the regenerating cell wall only occurs after the establishment of a structural glucan-chitin matrix around the plasma membrane of reverting protoplasts. Consequently, during the early stages of the regeneration process of protoplast walls, cell wall protein precursors are not retained into the nascent wall but are secreted into the culture medium. It follows that deposition of the fibrillar component (polysaccharide framework in the inner wall layer) and amorphous component (protein precursors in the outer wall layer) into the de novo cell wall is therefore desynchronized (5). The filasomes, located in the reverting site of the protoplasts, appear to be involved in the transport of cell wall components from the cytoplasm to the plasma membrane in association with secretory vesicles. Interestingly, actin plays a key role in (1) the initiation of cell wall regeneration, (2) the correct deposition of cell wall precursors, and (3) preservation of the proper shape of the regenerating protoplasts (see Note 2) (15,23).

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7. Discover new diagnostic/prognostic markers and/or therapeutic targets for human mycoses (14). 8. Increase the productivity and facilitate downstream processing of useful (recombinant or nonrecombinant) proteins extracellularly expressed in yeasts (16), because the use of regenerating protoplasts eliminates the physical barrier of the cell wall, which may limit their excretion. However, it is worth mentioning that this stratagem cannot be exploited for long-term production processes because of (1) the extreme fragility of yeast protoplasts, and (2) complete cell wall regeneration around the active protoplasts during long-term culture periods (see Note 3).

In this chapter, we describe a method for obtaining proteins secreted into the culture medium during the early stages of the regeneration process of protoplast walls under suitable conditions for subsequent proteomic analyses (10,13,14,17,18). We also provide a rapid and straightforward protocol for checking the degree of purity of the cell wall proteins excreted from yeast protoplasts in active cell wall regeneration. Proteins involved in cell wall construction (such as -1,3-glucosyltransferases, exoglucanases, glycosyl phosphatidylinositol (GPI)-proteins, and proteins with internal repeats [PIR], among others), heat shock proteins, glycolytic enzymes and other proteins have successfully been identified using the procedure presented here in combination with 2DE and MS (10,13,14,17). Given the nature of the protein sample, (multidimensional) liquid chromatography techniques in tandem with MS could also alternatively be applied. Although other methods to isolate and solubilize cell wall proteins from yeast species have been reported in the Chapter 19, the choice between these or the one outlined here, which each have advantages and disadvantages, will depend on the specific application.

2. Materials All solutions and buffers should be prepared with ultrapure water (doubledistilled, deionized water with a resistivity >18M/cm), and prechilled when working at 4°C. Growth media, solutions and buffers should be sterilized by autoclaving before use. Their labile components should be filter-sterilized separately and added to the other ingredients after autoclaving. 2.1. Collection of Proteins Secreted from Yeast Protoplasts in Active Cell Wall Regeneration 2.1.1. Preparation of Yeast Protoplasts 1. Yeast-Peptone-D-glucose (YPD) plates: 1% (w/v) yeast extract (Difco Laboratories, Detroit, MI), 2% (w/v) peptone (Difco), 2% (w/v) d-glucose, 2% (w/v) agar (Difco).

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2. YPD medium: 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) d-glucose. 3. Pretreatment buffer: 10 mM Tris-HCl, pH 9.0, 5 mM EDTA, 1% (v/v) -mercaptoethanol. 4. 1 M sorbitol solution: Dissolve 182.17 g of sorbitol in sufficient water to yield a final volume of 1 L. 5. Glusulase® (Du Pont; NEN Life Science Products, Boston, MA).

2.1.2. Active Cell Wall Regeneration of Yeast Protoplasts 1. Complete minimal (CM) medium: 0.17% (w/v) yeast nitrogen base (YNB) without amino acids or ammonium sulfate (Difco), 0.5% (w/v) (NH4 )2 SO4 , 2% (w/v) d-glucose, 0.13% (w/v) dropout powder (see Note 4). 2. Dropout powder: 2.5 g adenine, 1.2 g l-arginine, 6.0 g l-aspartic acid, 6.0 g lglutamic acid, 1.2 g l-histidine, 3.6 g l-leucine, 1.8 g l-lysine, 1.2 g l-methionine, 3.0 g l-phenylalanine, 22.5 g l-serine, 12.0 g l-threonine, 2.4 g l-tryptophan, 1.8 g l-tyrosine, 9.0 g l-valine, and 1.2 g uracil. 3. Lee medium (19): 0.02% (w/v) MgSO4 , 0.25% (w/v) K2 HPO4 , 0.5% (w/v) NaCl, 0.5% (w/v) (NH4 )2 SO4 , 0.05% (w/v) l-alanine, 0.13% (w/v) l-leucine, 0.1% (w/v) l-lysine, 0.01% (w/v) l-methionine, 0.007% (w/v) l-ornitine, 0.05% (w/v) l-phenylalanine, 0.05% (w/v) l-proline, 0.05% (w/v) l-threonine (see Note 5). After autoclaving, add 25 mL of 50% glucose solution and 2 mL of 0.1% biotin solution. 4. 50% glucose solution: Dissolve 12.5 g of d-glucose in a final volume of 25 mL water and sterilize using a 0.22-μm filter (Millipore, Bedford, MA). 5. 0.1% biotin solution: Dissolve 2 mg of biotin in a final volume of 2 mL water and sterilize using a 0.22-μm filter.

2.1.3. Recovery of Proteins Secreted from Regenerating Protoplasts 1. Phenylmethylsulfonyl fluoride (PMSF) stock solution: 0.1 M PMSF in isopropanol. Dissolve 174 mg of PMSF (Fluka, Chelmsford, MA) in a final volume of 10 mL isopropanol, and store at –20 ºC (see Note 6). PMSF should be handled with caution, because it is highly toxic. Weigh this hazardous chemical in a fume hood, and wear gloves, goggles and a mask. 2. Antipain stock solution: Dissolve 5 mg of antipain (Sigma, St. Louis, MO) in a final volume of 1 mL water, and store at –20 ºC (see Note 7). 3. Leupetin stock solution: Dissolve 5 mg of leupeptin (Sigma) in a final volume of 1 mL water, and store at –20 ºC (see Note 8). 4. Pepstatin stock solution: Dissolve 2.5 mg of pepstatin (Sigma) in a final volume of 1 mL methanol, and store at –20 ºC (see Note 9). 5. Protease inhibitor mix: 0.1 mM PMSF, 2 μg/mL antipain, 2 μg/mL leupeptin, and 1 μg/mL pepstatin (i.e., for 450 mL of protein solution, add 450 μL of PMSF stock solution and 180 μL each of antipain, leupeptin and pepstatin stock solutions). Mix just before use. 6. Filtration unit (Millipore). 7. 0.22-μm pore-size nitrocellulose filter (Millipore).

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2.1.4. Concentration of Proteins Secreted from Regenerating Protoplasts 1. Ultrafiltration apparatus (300-mL stirred ultrafiltration cell system; Amicon; Beverly, MA). 2. YM-10 Diaflo® ultrafiltration membrane (pre-equilibrated 10,000-Da pore-size ultrafilter; Amicon). 3. Magnetic stirrer. 4. Oxygen-free nitrogen with pressure regulator (see Note 10). 5. Membrane wash solution: 1–2 M NaCl. 6. Membrane store solution: 10% ethanol. 7. Lyophilizer.

2.2. Assay of Cell Wall Protein Purity: Alkaline Phosphatase Assay 1. Lysis buffer: 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM dithiothreitol (DTT), 0.5 mM PMSF and 5 μg/mL each of antipain, leupeptin and pepstatin (see Notes 6–9). 2. NaCl-glycine buffer (alkaline buffer): 0.1 M NaCl, 0.1 M glycine, pH 9.7 (adjusted with 1 N NaOH). 3. Stock p-nitrophenol (PNP) solution: 8 mM PNP (Sigma) in NaCl-glycine buffer. 4. Substrate solution: 20 mM disodium p-nitrophenyl phosphate (PNPP; Sigma) in NaCl-glycine buffer. 5. Stop solution: 0.05 M NaOH. 6. Spectrophotometer.

3. Methods 3.1. Collection of Proteins Secreted from Yeast Protoplasts in Active Cell Wall Regeneration The flowchart in Fig. 2 summarizes the different steps in the collection of proteins excreted into the osmotically stabilized liquid nutrient medium during the first stages of the regeneration process of protoplast walls. These involve (1) the complete elimination of the yeast cell wall by controlled enzymatic digestion (i.e., protoplast formation), (2) active regeneration of the cell wall and ensuing secretion of wall protein precursors into the growth medium, (3) recovery of the culture medium with secreted proteins by centrifugation and filtration, and (4) concentration of secreted proteins by ultrafiltration and lyophylization. This method to isolate and solubilize protein precursors of the yeast cell walls is taken from the published protocols in Saccharomyces cerevisiae and Candida albicans by Pardo et al. (10) and Pitarch et al. (17), respectively, and is based on earlier methodology developed by Elorza et al. (20). Although this procedure has been used successfully and reproducibly on

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Fig. 2. Flowchart of basic steps in the collection of proteins secreted from yeast protoplasts in active cell wall regeneration into the osmotically stabilized liquid nutrient medium.

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these yeast species for subsequent proteomic analyses (10,13,14,17), it may be necessary to adjust it for other yeast species or even for filamentous fungi. 3.1.1. Preparation of Yeast Protoplasts (see Note 11) 1. Grow yeast cells on a YPD plate (stock maintenance medium) at 28–30 ºC for 2 d. Inoculate 50 mL of YPD medium in a 250-mL flask (see Note 12) with a single colony from the YPD plate, and grow overnight at 28–30 ºC in a shaking incubator (200 rpm). 2. Use this 50-mL preculture to inoculate two 2-L flasks containing 500 mL fresh YPD medium each, and grow at 28–30 ºC with vigorous rotary shaking (200 rpm) until the culture reaches mid-log phase growth (OD600nm = 4; see Note 13). 3. Harvest the yeast cells by centrifugation at 4,500g for 5 min and discard the supernatant. 4. Resuspend the cell pellet in 150 mL water, and centrifuge 5 min at 4,500g. Decant the supernatant. 5. Gently resuspend the yeast cells in 100 mL of pretreatment buffer to a density of 1–2 × 109 cells/mL, and incubate at 28 ºC with gentle rotary shaking (80 rpm) for 30 min (see Note 14). Centrifuge 10 min at 600g, and discard the supernatant. 6. Gently resuspend the cell pellet in 150 mL of a 1 M sorbitol solution (see Note 15). Centrifuge 10 min at 600g to harvest cells, and discard the supernatant. 7. Gently resuspend the cell pellet in a 1 M sorbitol solution to a density of 5 × 108 cells/mL, and add 30 μL/mL ice-cold Glusulase® (see Note 16). 8. Incubate cells with very gentle shaking (80 rpm; see Note 17) at 28 ºC until more than 90–95% of them are protoplasts (∼45 min to 1 h). Monitor the degree of protoplast formation with a phase-contrast microscope (see Note 18). 9. Harvest protoplasts by very gentle centrifugation at 600g for 15 min. Decant the supernatant carefully (see Note 19). 10. Gently wash the protoplast pellet with 150 mL of a 1 M sorbitol solution (see Note 20). Centrifuge 15 min at 600g and decant the supernatant carefully. 11. Repeat this step two more times to eliminate any trace of Glusulase® (see Note 21). 12. Remove a small aliquot of the protoplast preparation for use in Subheading 3.2 to evaluate enzymatic activity of alkaline phosphatase (see Note 22).

3.1.2. Active Cell Wall Regeneration of Yeast Protoplasts (see Notes 11 and 17) 1. Gently resuspend the protoplasts in the regenerating buffer supplemented with 1 M sorbitol to a density of 3 × 108 cells/mL (see Notes 20 and 23). 2. Incubate protoplasts with gentle rotary shaking (80 rpm) at 28 ºC for 2 h to induce their active cell wall regeneration.

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3.1.3. Recovery of Proteins Secreted from Regenerating Protoplasts (see Notes 17 and 24) 1. Centrifuge the regenerating protoplasts at 600g for 15–20 min at 4 ºC, and collect the supernatant carefully (see Note 19 and 25). 2. Add the protease inhibitor mix. 3. Filter the supernatant through a 0.22-μm pore-size nitrocellulose filter without any external device in ice at 4 ºC (see Note 26). 4. Remove a small aliquot of cell-free culture filtrate to further monitor the cell lysis by quantitative determination of alkaline phosphatase (see Subheading 3.2).

3.1.4. Concentration of Proteins Secreted from Regenerating Protoplasts (see Note 24) 1. Wash an YM-10 Diaflo® ultrafiltration membrane by floating it skin (glossy) side down in a beaker of water for 1 h (see Note 27). Change water at least three times. 2. Concentrate the cell-free culture filtrate by ultrafiltration using the washed YM-10 ultrafilter (see Fig. 3A and Notes 10 and 28). 3. Dilute with 300 mL water, and concentrate again. Repeat this step three more times to eliminate any trace of sorbitol (see Note 29). 4. Remove the ultraconcentrated protein solution (molecular weight fraction above 10,000 Da) from the ultrafiltration cell (see Fig. 3B and Note 30). 5. Quick-freeze the ultraconcentrated protein solution at –80 ºC (see Note 31), and concentrate it by lyophilization (see Note 32). 6. Resuspend the freeze-dried protein sample in a small volume of water (see Note 33), and quantify the protein content by standard protein determination methods.

3.2. Assay of Cell Wall Protein Purity: Alkaline Phosphatase Assay (see Note 34) The method given below outlines an easy and prompt screening procedure to check the efficiency, purity, and quality of the collection of proteins secreted from yeast protoplasts in active cell wall regeneration before performing further proteomic analyses and establishing any interpretation of the results. This is based on measurement of enzymatic activity of alkaline phosphatase, which is exclusive to the cytosol in yeasts and thus exploited as a marker of cytosolic contamination. This intracellular enzyme catalyzes the cleavage of phosphate ester bonds from many compounds (including the chromogenic phosphatase substrate used in this protocol, p-nitrophenylphosphate; PNPP) under alkaline conditions and an optimum temperature of 37 ºC. If there is protoplast lysis during sample preparation and, therefore, cytosolic contamination (containing alkaline phosphatase, among other intracellular proteins) into the sample of

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Fig. 3. Schematic representation of a stirred ultrafiltration cell on a magnetic stirrer. Pressurized stirred ultrafiltration cells can efficiently be used to simultaneously concentrate and desalt samples of dilute proteins (from initial volumes of 3 mL–2 L to final concentrate volumes of 50 μL–60 mL). In the procedure described here, the cell-free culture filtrate is forced through an YM-10 Diaflo® ultrafilter, located on a polymer grid at the bottom of the cell (see Note 28). Protein sample is separated into two groups according to molecular weight and size (see Note 27). The ultraconcentrate (molecular weight fraction above 10,000 Da) is subjected to lyophilization or centrifugal microconcentration (see Note 32) before carrying out further proteomic analyses. The protein content of the ultrafiltrate (molecular weight fraction below 10,000 Da) can also be quantified by standard protein determination methods to evaluate the final protein recovery.

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wall protein precursors, then alkaline phosphatase will hydrolyze PNPP (which is colorless), releasing p-nitrophenol (PNP, which is yellow-colored at alkaline pH but colorless at acidic pH values) and inorganic phosphate (see Fig. 4). Accordingly, only protein solutions with intracellular contamination will turn yellow following the addition of PNPP. The enzyme activity of alkaline phosphatase can consequently be detected and quantified by spectrophotometry.

Fig. 4. Biochemical principle of the alkaline phosphatase assay. The pnitrophenylphosphate (PNPP) is a chromogenic substrate for several phosphatases, such as acid phosphatases and alkaline phosphatases. Alkaline phosphatase hydrolyzes this artificial substrate, which is colorless, at alkaline pH values (pH = 9.7) and 37°C to form p-nitrophenol (PNP), which is yellow-colored in basic solutions and can be measured at 410 nm on a spectrophotometer. The intensity of the yellow color correlates with the amount of PNP issuing from the hydrolysis of PNPP catalyzed by phosphatase alkaline, according to Beer’s law. In contrast, acid phosphatase hydrolyzes PNPP under acidic conditions (pH = 4.8) and 37°C leading to PNP, which is colorless at acidic pH values.

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The present method is extremely sensitive, because very small amounts of alkaline phosphatase generate sufficient PNP after an adequate incubation period, bringing a measurable color about. This protocol is modified from that of Cabib et al. (21), which was developed for rapid checking of colonies with lysed yeast cells (embedded into a gelled medium supplemented with PNPP at pH 9.7). These acquired a yellow color on and around colonies as a result of the release of alkaline phosphatase. Interestingly, the present assay is not affected by the acid phosphatases, which are located in the outermost and innermost cell wall layers of the yeast cells (1) and, therefore, detected in intact cells (see Fig. 4) . This is because of the fact that these extracellular enzymes are only able to hydrolyze PNPP under acidic conditions (optimum pH = 4.8) and the pH of the reaction buffer is highly alkaline (pH = 9.7). 1. Resuspend the aliquot of protoplasts (see Subheadings 3.1.3 and Note 22) in 1 mL of ice-cold lysis buffer. Vortex for 1 min, and cool on ice for 1–2 min. Repeat this step until complete cell breakage (monitored with a phase-contrast microscope). Remove cell debris from the homogenate by centrifugation at 12,500g for 10 min at 4 ºC. Collect the supernatant and use it as a source of alkaline phosphatase for the positive control (see Note 35). 2. Use known amounts of the reaction product PNP in NaCl-glycine buffer (10–110 μg/mL) to obtain a standard curve. Mix thoroughly by careful vortexing. Measure and record the absorbance at 410 nm in the spectrophotometer. Use the same plastic cuvet for all samples, starting with the most dilute sample. 3. Include controls of (1) NaCl-glycine buffer alone (buffer control), (2) cell-free culture filtrate alone (enzyme control), (3) substrate without cell-free culture filtrate (substrate control), and (4) substrate with protoplast lysate (positive control) as indicated in Fig. 5 to determine superfluous absorbance either from other compounds in buffer and cell-free culture filtrate, or from substrate breakdown (see Note 36). 4. Place 500 μL of cell-free culture filtrate of products secreted from regenerating protoplasts in a 1.5-mL Eppendorf tube (see Subheadings 3.1.3 and Note 36), and add 500 μL of substrate solution (see Note 36). 5. Vortex carefully and incubate in the dark at 37 ºC for 30 min. Protect from bright light. 6. Stop the enzyme activity with 400 μL of stop solution. Mix carefully and cool the tubes to room temperature (25–35ºC). 7. Measure and record the absorbance at 410 nm in the spectrophotometer. Use the same plastic cuvet for all tubes. 8. Determine enzyme activities in all tubes using the standard curve (10–110 μg p-nitrophenol/mL). The units of enzyme (see Note 37) are calculated as: U.E. = 14.38 × c/t, where c is the PNP concentration in μg/mL and t is the reaction time in min.

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Fig. 5. Schematic diagram of the basic steps in the alkaline phosphatase assay.

4. Notes 1. In all yeast species studied so far, protoplasts embedded into gelled nutrient media are always able to regenerate a complete cell wall and revert to normal reproducing cells (5). In contrast, the cell wall regeneration of reverting yeast protoplasts grown in liquid nutrient media is (1) incomplete in Saccharomyces cerevisiae and other budding yeasts, bringing about cells with aberrant walls (albeit with all cell wall components) incapable of dividing and inducing normal morphogenesis (5,12), but (2) complete in Candida albicans (4,22), Schizosaccharomyces pombe (23,24), Nadsonia elongata (5) and Endomycopsis fibuliger (5), resulting in normal growing cells. Accordingly, certain physical

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2.

3.

4.

5. 6.

7.

8.

9.

10.

11. 12. 13.

14.

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factors can actively be involved in the cell wall construction and protoplast reversion (5). The morphology of the de novo cell wall formed on the protoplast surface follows that of the protoplast (i.e., spherical shape) (5). On the contrary, the shape of new cells originated from the walled protoplasts is at least partly determined by the cell wall (i.e., oval shape; see Fig. 1). The complete reversion of S. pombe, N. elongata and C. albicans protoplasts to normal cell walls and growing cells is attained after 12, 20, and 24 h, respectively, of incubation in osmotically stabilized liquid nutrient media (4,5,23). To promote optimal regeneration of protoplast cell walls, it is convenient to use a 10× dropout powder solution that has been autoclaved separately or filter sterilized, and added to the remaining ingredients after autoclaving. To avoid precipitation of MgSO4 and K2 HPO4 , these should be autoclaved separately or filter sterilized, and added to the other ingredients after autoclaving. The PMSF can also be solubilized in ethanol, methanol and 1,2-propanediol; and is unstable in aqueous solution. PMSF inhibits serine proteases (e.g., trypsin, chymotrypsin, and thrombin) and thiolproteases (e.g., papain). It is added to reduce possible proteolytic processes (see Table 1). Antipain is also soluble in methanol and dimethylsulfoxide (DMSO). It inhibits papain and trypsin and, to a lesser extent, plasmin. It is added to reduce possible proteolytic processes (see Table 1). The inhibition specificity of leupeptin is of broad spectrum. It inhibits serine and thiol-proteases. It is added to reduce possible proteolytic processes (see Table 1). Pepstatin is insoluble in water. It inhibits acid proteases (e.g., pepsin, chymosin, cathepsin D and renin, among others). It is added to reduce possible proteolytic processes (see Table 1). It is important not to use compressed air because it can bring about (1) large pH changes as a result of carbon dioxide dissolution, and/or (2) oxidations in sensitive protein solutions. Perform all procedures from this subheading under sterile conditions. Use sterile centrifuge bottles. Liquid cultures should be grown in a flask that is at least 4–5 times larger than the culture volume. The cell density of inoculum and incubation time should be adjusted according to the yeast strain. It is important that the yeast culture is in mid-log phase growth (∼ OD600nm = 4) because (1) yeast cells are more susceptible to wall lytic enzymes (Glusulase® treatment) for complete protoplasting than those in stationary phase growth and (2) there is sufficient biomass accumulation. This step is critical for a successful protoplast preparation because mercaptoethanol pretreatment facilitates the subsequent action of wall lytic enzymes (Glusulase® treatment) by (1) breaking disulfide bonds of cell wall proteins and (2) slightly disorganizing the cell wall. In fact, recent studies have demonstrated that a better protoplasting effect is attained in

Serine proteases Thiolproteases

Acid proteases

Leupeptin

Pepstatin

0.7–1 μg/mL

b

Reversible by dithiothreitol (DTT) treatment. Serine protease. c Thiolprotease. d Dimethylsulfoxide. e Sit overnight.

a

2–50 μg/mL

Trypsinb Papainc

Antipain

0.5–2 μg/mL

17–174 μg/mL (0.1–1 mM)

Serine proteases Thiolproteasesa

PMSF

Effective concentrations

Specificity

Inhibitor

Table 1 Protease inhibitors

2.5 mg/mL in methanol

5 mg/mL in water

5 mg/mL in water

17 mg/mL in isopropanol

Stock solutions

Methanol Ethanole

Isopropanol Ethanol Methanol 1, 2-propanediol Water Methanol DMSOd Water

Soluble in

Stable ∼6 months at –20 ºC Stable ∼1 week at +4 ºC Stable ∼1 week at +4 ºC

Stable ∼1 month at –20 ºC

Notes

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16.

17.

18.

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-mercaptoethanol-pretreated yeast cells than in nonpretreated cells (25). Centrifugation for isolation of -mercaptoethanol-treated yeast cells should be carried out with the brake in 1.5. 1 M sorbitol is used as an osmotic stabilizer. In general, sugars and sugar alcohols (e.g., sorbitol, mannitol) are consistently used for yeast protoplast formation and reversion, whereas MgSO4 and KCl are commonly chosen for preparation and regeneration of protoplasts from filamentous fungi. Viable protoplasts can be achieved by controlled autolysis of the yeast cell wall using snail or microbial lytic enzymes (see Table 2) (25). Glusulase® is a preparation of the intestinal juice of the Roman garden snail Helix pomatia, and consists of a mixture of lytic enzymes (-glucuronidase, sulfatase, and cellulase). It has proven particularly useful for obtaining protoplasts (i.e., for completely digesting yeast cell walls) in nearly all yeast species (5). Sterilized tweezers should be used to take the lid off the Glusulase® bottle. It is important not to do it with the hands. Gently mix Glusulase® with cell suspension (without shaking it). Perform all procedures from this point on with very gentle shaking. It is very important to handle the protoplasts gently in this protocol, because yeast protoplasts are extremely fragile. This can prevent further intracellular contamination into the sample of proteins secreted from actively reverting protoplasts. In general, 90–95% of protoplasts should be routinely achieved. The incubation time and amount of Glusulase® needed to attain protoplasts (i.e., complete yeast cell wall lysis) can vary with yeast strain and growth stage and may therefore be necessary to adjust the incubation time and Glusulase® amount given in the protocol. The degree of protoplast formation is assessed under a phase-contrast microscope by: a. Counting spherical cells. b. Observing cell lysis in hypotonic solution (e.g., after the addition of water; see Fig. 6). The number of osmotically-resistant cells (non-protoplasted cells) is determined by diluting an aliquot of the cell suspension after and before Glusulase® treatment in water and plating on YPD plates (and growing at 28–30 ºC). The ratio of CFUs after to before treatment is then calculated to estimate the efficiency of protoplast formation.

19. Perform all centrifugations for protoplast isolation with the brakes off. It is important to decant the supernatant carefully, because the protoplast pellet is less compact than the preceding cell pellets. 20. Protoplast resuspension is difficult and sticky. For this reason, a small volume should first be used to resuspend the protoplasts by gently swirling liquid across the surface of the pellet. Then add more solution until reaching the correct final volume. 21. It is important to wash the protoplasts several times (before regenerating protoplast walls) to remove sulfatases, and on the whole any enzymatic activity,

b

a

Gut juice (digestive enzymes) Gut juice (digestive enzymes) Culture fluid

Achatina achatina (the giant African snail) Arthrobacter luteus (bacterium)

Location

Helix pomatia (the Roman garden snail)

Source

Very expensive for use in industrial processes. Trace.

Microbial enzymes a

Snail enzymes

Types

-glucuronidase endo--glucanase arylsulfatase -1,3-glucan laminaripentaohydrolase – -1,3-glucanase – protease – mannase – amylaseb – xylanaseb – phosphataseb

– – – –

– -glucuronidase – sulfatase – cellulase

Composition

Zymolyase 20T® – MP Biomedicals, Aurora, Ohio, US – Seikagaku Corporation, Tokyo, Japan – Miles Laboratories, Elkhart, Ill., US

(10, 14, 26)

Glusulase® - Du Pont; NEN Life Science Products, Boston, MA, US Not available

(20, 28)

(25, 27)

References

Commercial name

Table 2 Main preparations of lytic enzymes used for yeast cell wall digestion and subsequent production of viable protoplasts from yeast cells

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Fig. 6. Evaluation of the efficiency of protoplast formation. When an osmotically stabilized protoplast/cell suspension is diluted with water, the extracellular medium becomes hypotonic to cytosol (i.e., the free water concentration is greater outside the cell). This results in a passive movement of free water from extracellular medium to cytoplasm (an influx of free water). Consequently, protoplasts gain water, swell and burst, whereas nonprotoplasted cells do not undergo cytolysis but an increase in turgor pressure because of the presence of the cell wall (see Note 18). In isotonic solutions, the movement of water into and out of the cell maintains equilibrium. present in the Glusulase® preparation that may modify the protein precursors secreted into the medium. 22. The protoplast preparation can be lysed to release intracellular proteins and used as a source of alkaline phosphatase (positive control; see Subheading 3.2). 23. Lee medium is reliably used as a regenerating buffer for Candida spp. (9,17), whereas complete minimal (CM) medium is preferred for Saccharomyces spp. (10). These regeneration media are chosen because they are rich and chemically

260

24.

25.

26.

27.

28.

29. 30.

31. 32. 33.

34.

Pitarch et al. defined media. Hence, there are no substances in their composition that interfere in the subsequent proteomic analyses of the secreted proteins. YPD medium is a complex medium, and cannot be used because it contains proteins (from the yeast extract), which could result in misidentifications. Perform all procedures from this point on at 4 ºC (in a cool room at 4 ºC) with precooled solutions, reagents and apparatus to avoid undesirable proteolytic activity. It is probable that the protoplast pellet is resuspendend when there is a small volume of the supernatant left before being carefully collected. If this happens, then centrifuge once more at 600 g for 20 min, and gently collect the supernatant again. If there is not enough time to finish the entire protocol in one laboratory period, the procedure can be stopped after filling the filtration unit with the centrifuged medium (supernatant) and placing it in a container with ice in a cool room at 4 ºC. Do not use any external device, because this may facilitate protoplast lysis and subsequent intracellular contamination in the sample of secreted proteins. The YM-10 Diaflo® ultrafiltration membrane had a pore size of 10,000 Da to yield molecular weight fractions above and below 10,000 Da. It should be handled carefully and only at the edge. The washing step is important to remove preservative substances (e.g., sodium azide). Place the YM-10 membrane in an ultrafiltration cell, skin (glossy) side toward cell-free culture filtrate, and then fill the cell with the culture filtrate. Place it on a magnetic stirrer, and connect the inlet line to a regulated nitrogen pressure source (see Note 10 and Fig. 3A). Pressurize the cell and pressure-check following the supplier’s instructions. Turn on the stirrer, and adjust the stirring rate. When the ultrafiltration is accomplished, depressurize and continue stirring for a few minutes to increase protein recovery (see Fig. 3B). It is essential to remove any trace of sorbitol because this interferes in the protein resolution of subsequent proteomic analyses. The ultrafiltration membrane should be washed with 1–2 mL of water to elute proteins potentially sticking to it. Add this volume to the ultraconcentrate. To reuse the ultrafilter, rinse it with 1–2 M NaCl, and then flush it through with water. Store the ultrafilter in 10% ethanol at 4 ºC. The ultraconcentrate can also be frozen quickly into liquid nitrogen. Centrifugal microconcentration techniques can alternatively be used to reconcentrate the ultraconcentrated protein sample. The protein sample can be used directly in two-dimensional gel electrophoresis (see Fig. 7) (10,13,14,17). Alternatively, freeze-dried protein preparation can be resuspended in a small volume of buffer suitable for the subsequent proteomic analyses. This assay should be handled throughout with caution, because p-nitrophenol is a skin irritant. Wear gloves and a laboratory coat. Duplicates of each tube should be performed.

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Fig. 7. Proteomics of proteins secreted from C. albicans protoplasts in active cell wall regeneration.

35. The clarified supernatant (protoplast lysate) can be stored at –80 ºC and used in future assays. 36. This assay can also be carried out on a microtiter plate, using 100 μL, rather 500 μL, of each component. 37. Although substantial enzyme activity is found in the absence of Mg2+ ions, it is convenient to use 1.5 mM MgCl2 or 1 mM MgSO4 in the substrate solution, because these divalent cations stimulate alkaline phosphatase activity. 38. One unit of enzyme (U.E.) is defined as the amount of enzyme that will produce 1 nmol of p-nitrophenol per minute.

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Acknowledgments We thank the Merck, Sharp & Dohme (MSD) Special Chair in Genomics and Proteomics, European Community (STREP LSHB-CT-2004-511952), Comunidad de Madrid (S-SAL-0246/2006) and Comisión Interministerial de Ciencia y Tecnología (BIO-2003-00030 and BIO-2006-01989) for financial support of our laboratory.

References 1. Chaffin, W. L., Lopez-Ribot, J. L., Casanova, M., Gozalbo, D. and Martinez, J. P. (1998) Cell wall and secreted proteins of Candida albicans: identification, function, and expression. Microbiol Mol.Biol Rev. 62, 130–80. 2. Klis, F. M., Boorsma, A., and de Groot, P. W. (2006) Cell wall construction in Saccharomyces cerevisiae. Yeast 23, 185–02. 3. Osumi, M. (1998) The ultrastructure of yeast: cell wall structure and formation. Micron. 29, 207–33. 4. Nishiyama, Y., Aoki, Y., and Yamaguchi, H. (1995) Morphological aspects of cell wall formation during protoplast regeneration in Candida albicans. J.Electron Microsc. 44, 72–8. 5. Necas, O. (1971) Cell wall synthesis in yeast protoplasts. Bacteriol.Rev. 35, 149–70. 6. Takagi, T., Ishijima, S. A., Ochi, H., and Osumi, M., (2003) Ultrastructure and behavior of actin cytoskeleton during cell wall formation in the fission yeast Schizosaccharomyces pombe. J.Electron Microsc. 52, 161–74. 7. Rico, H., Carrillo, C., Aguado, C., Mormeneo, S., and Sentandreu, R. (1997) Initial steps of wall protoplast regeneration in Candida albicans. Res.Microbiol. 148, 593–603. 8. Kapteyn, J. C., Dijkgraaf, G. J., Montijn, R. C., and Klis, F. M. (1995) Glucosylation of cell wall proteins in regenerating spheroplasts of Candida albicans. FEMS Microbiol Lett. 128, 271–77. 9. Elorza, M. V., Marcilla, A., Sanjuan, R., Mormeneo, S., and Sentandreu, R. (1994) Incorporation of specific wall proteins during yeast and mycelial protoplast regeneration in Candida albicans. Arch.Microbiol. 161, 145–51. 10. Pardo, M., Monteoliva, L., Pla, J., Sanchez, M., Gil, C., and Nombela, C. (1999) Two-dimensional analysis of proteins secreted by Saccharomyces cerevisiae regenerating protoplasts: a novel approach to study the cell wall. Yeast 15, 459–72. 11. Pitarch, A., Sanchez, M., Nombela, C., and Gil, C. (2002) Sequential fractionation and two-dimensional gel analysis unravels the complexity of the dimorphic fungus Candida albicans cell wall proteome. Mol.Cell Proteomics 1, 967–82. 12. Klis, F. M. (1994) Review: cell wall assembly in yeast. Yeast, 10, 851–69. 13. Pardo, M., Ward, M., Bains, S., et al. (2000) A proteomic approach for the study of Saccharomyces cerevisiae cell wall biogenesis. Electrophoresis 21, 3396–3410. 14. Pitarch, A., Jimenez, A., Nombela, C., and Gil, C. (2006) Decoding serological response to Candida cell wall immunome into novel diagnostic, prognostic, and

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15.

16.

17.

18.

19.

20.

21. 22.

23.

24.

25.

26. 27.

28.

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therapeutic candidates for systemic candidiasis by proteomic and bioinformatic analyses. Mol.Cell Proteomics 5, 79–96. Kobori, H., Yamada, N., Taki, A., and Osumi, M. (1989) Actin is associated with the formation of the cell wall in reverting protoplasts of the fission yeast Schizosaccharomyces pombe. J.Cell Sci. 94, 635–46. Tanaka, H., Kamogawa, T, Aoyagi, H., Kato, I. and Nakajima, R. (2000) Invertase production by Saccharomyces cerevisiae protoplasts immobilized in strontium alginate gel beads. J. Biosci. Bioeng. 89, 498–500. Pitarch, A., Pardo, M., Jimenez, A., et al. (1999) Two-dimensional gel electrophoresis as analytical tool for identifying Candida albicans immunogenic proteins. Electrophoresis 20, 1001–10. Pitarch, A., Nombela, C., and Gil, C. (2006) Candida albicans biology and pathogenicity: Insights from proteomics, in Microbial Proteomics: Functional Biology of Whole Organisms (Humphery-Smith,I. and Hecker,M., eds), Wiley-Vch, Hoboken, NJ, pp. 285–330. Lee, K. L., Buckley, H. R., and Campbell, C. C. (1975) An amino acid liquid synthetic medium for the development of mycelial and yeast forms of Candida albicans. Sabouraudia. 13, 148–53. Elorza, M. V., Rico, H., Gozalbo, D., and Sentandreu, R. (1983) Cell wall composition and protoplast regeneration in Candida albicans. Antonie Van Leeuwenhoek 49, 457–69. Cabib, E. and Duran, A. (1975) Simple and sensitive procedure for screening yeast mutants that lyse at nonpermissive temperatures. J.Bacteriol. 124, 1604–06. Murgui, A., Elorza, M. V., and Sentandreu, R. (1986) Tunicamycin and papulacandin B inhibit incorporation of specific mannoproteins into the wall of Candida albicans regenerating protoplasts. Biochim.Biophys.Acta 884, 550–8. Osumi, M., Yamada, N., Kobori, H., et al. (1989) Cell wall formation in regenerating protoplasts of Schizosaccharomyces pombe: study by high resolution, low voltage scanning electron microscopy. J.Electron Microsc. 38, 457–68. Osumi, M., Sato, M., Ishijima, S. A., Konomi, M., Takagi, T., and Yaguchi, H. (1998) Dynamics of cell wall formation in fission yeast, Schizosaccharomyces pombe. Fungal.Genet.Biol. 24,178–206. Ezeronye, O. U. and Okerentugba, P. O. (2001) Optimum conditions for yeast protoplast release and regeneration in Saccharomyces cerevisiae and Candida tropicalis using gut enzymes of the giant African snail Achatina achatina. Lett.Appl.Microbiol. 32, 190–93. Eddy, A. A. and Williamson, D. H. (1957) A method of isolating protoplasts from yeasts. Nature 179, 1252–53. Agogbua, S. O., Anosike, E. O and Ugochukwu, E. N. (1978) Partial purification and some properties of arylsulphatases from the gut of the giant African snail Achatina achatina. Comp. Biochem. Phys. 59B, 169–73. Kaneko, T., Kitamura, K. and Yamamoto, Y. (1973) Susceptibilities of yeast to yeast cell lytic enzyme of Arthrobacter luteus. Agric. Biol. Chem. 37, 2295–2302.

21 Sample Preparation Procedure for Cellular Fungi Alois Harder

Summary A crucial step in quantitative proteomics is an artefact free and reproducible sample preparation protocol, which has to be adapted and optimized to nearly all types of cells. Here we provide a sample preparation method for quantitative proteomics of cellular fungi. Two different protein extraction methods were compared with focus on reproducibility, minimized proteolytic degradation and protein losses during the sample preparation. In the first preparation the cells were lysed by sonication followed by protein solubilization in “standard” lysis buffer. The second preparation was performed with a SDSpresolubilization step followed by sonication and further boiling, before diluting the sample with lysis buffer. We have shown that the sample preparation for cellular fungi is performed with maximum protein solubilization, higher reproducibility and a reduced proteolytic activity by including a SDS-presolubilization step in the sample preparation protocol.

Key Words: 2D-electrophoresis; fungi; proteolytic activity; sample preparation; yeast.

1. Introduction Fungal cells have nowadays found their way in nearly all areas of biochemistry and pharmacy. Famous examples are active medicaments like PerenterolTM with Saccharomyces boulardii as the active ingredient (1,2), yeast host cell systems like the vaccine production cycle or diverse secondary metabolite discovering systems like the Novobiocin synthesis to name only few (3,4). To push those applications or detect new fungal targets or metabolites, a solid basis of information in cell growth, cell cycle, gene- and protein data are essential. Whereas cell growth and cycle can be researched in conceivable From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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time, gene and protein data are missing for the majority of the fungi. The finished and currently running fungal genome sequencing projects are mainly concentrating on model- or pathogen strains: Saccharomyces cerevisiae, Ashbya gossypii, Aspergillus fumigatus, Candida parapsilosis, Candida dubliniensis, Pneumocystis carinii, Phytophthora infestans, and Schizosaccharomyces pombe have been sequenced or will be finished soon and are available for the community (5,6). For identification of fungal proteins, genome data of the organism to be researched has to be available. Although the fungal protein databases are growing, the listed open reading frames are far from their complete and correct functional characterization. However, with the sequenced fungal genomes, the available protein data and the homology – search programs with a solid foundation for further proteome analysis where eumycotic cells are present. Fungal cells are easy to cultivate, to manipulate, and in most cases are non-toxic, but the preparation of a robust and artefactfree protein extract for a proteomics approach is a challenge. Major problems in fungal proteomics result from the cells strong proteolytic activity. Most of the fungal proteases, intensively researched in the eumycotic model organism Saccharomyces cerevisiae, are located in the cytosol and the vacuole. Few proteases are located in the membrane, the endoplasmic reticulum, the mitochondria, and the Golgi complex (7,8). These include endoproteinases, carboxypeptidases, aminopeptidases, and dipeptidyl-aminopeptinase. By activation of these proteases protein modifications will occur and can rule out quantitative proteomics research (9–10,11). Of the many cellular proteases, the lumenal vacuolar proteases comprise the major source of problems for protein analysis and proteome research. Endoproteinase A and B (PrA and PrB), carboxypeptidases Y and S, aminopeptidase I (LAP IV), and the aminopeptidase yscCo (ApCo) are found soluble in the vacuole (12,13). Polypeptide inhibitors of these vacuolar proteases are found in the cytosol (e.g., inhibitor for PrB is IB ; inhibitor for yscCo is yscCoF ). By preparing the crude extract all cell compartments are broken up, meaning that corresponding protease inhibitors will bind to their substrate by forming an inactive complex—in this state proteolytic activity is inhibited. As reported from Jones et al. the fungal proteases PrA and PrB, can be activated out of the crude extract by lowering the pH to a level of 4–5. The reason for this activation might be the hydrolysis of the polypeptide inhibitors (7,14). Further research on protease deficiency mutants showed that the addition of denaturating agents activates proteolysis by removing the corresponding inhibitor from the protease molecule. Due to the exigency in proteomics research, for working in strict

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denaturation conditions, the necessity of a “non standard” sample preparation procedure for fungal cells is obvious. The fungal sample preparation for proteomics research has to • (BL)be carried out in strong denaturating conditions for its stability during the 2D - electrophoretic separation • display robustness to guarantee reproducibility by repeating the experiments • solubilize low abundant protein as well as high and low molecular weight proteins in all pH ranges • effectively inactivate the strong proteolytic activity of fungal cells

2. Materials 2.1. Cell Culture 1. Standard aerobic incubator with an inbuilt horizontal rotary shaker. 300 mL Erlenmeyer flasks with silicone plugs are used for aerobic cell cultivation. 2. Horizontal rotary shaker (e.g., HS 260, IKA). 3. Incubation mask (e.g., Certomat H/HK). 4. S. cerevisiae haploid wild type strain N318C (“Deutsche Sammlung für Mikroben und Zellinien”, DSMZ Braunschweig, Germany). 5. Defined synthetic YNB medium (Difco Labs, San Francisco CA, USA) is dissolved to a concentration of 6.7 g/L in distilled water (<18.4 μS). 6. The medium then is supplemented with 20 g/L (w/v) glucose and buffered with sodium hydroxide (6 g/L, w/v) / succinic acid (10 g/L, w/v) to pH 5.8. 7. For a control smear a Sabouraud agar plate is used.

2.2. S - 35 In Vitro Cell Labeling 1. S 35 - methionine from GE Healthcare (SJQ0079-2.5MCi). 2. 2 × 4 cm cut filter paper (GE Healthcare, 80-110619) is used for drying and counting the labelled protein. 3. Scintillation cups (12 mL), e.g., from Sigma. 4. Liquid scintillation counter (e.g., Beckman LS 6000 IC / 1801). For detailed information in working prescriptions with radioactivity, calibrating of the scintillation counter, waste disposal, radioactive sample storage etc. please refer to reference (16).

2.3. Cell Disruption and Protein Solubilization 1. Lysis buffer: 7M urea, 2M thiourea, 4% CHAPS (w/v), 1% DTT (w/v), 0.5% Pharmalytes, pH 3–10 (v/v) and 10 mM Pefabloc protease inhibitor. 2. Presolubilization SDS-buffer (95°C): 1% (w/v) SDS, 100 mM Tris-HCl, pH 7.0. 3. Sonifier (Bandelin 60W, HD 2070 UW).

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2.4. Scintillation Counting and Protein Assay 1. For S35-labelled cells, a scintillation counting is performed: 4.5 mL POPOP solution is required for one vial; 2 × 4 cm cut filter paper is used for the counting (see Note 1). As a calibration standard a C12 isotope (GE Healthcare) is applied. 2. Lowry protein assay kit for SDS containing samples. 3. The Bradford protein assay kit (Bio-Rad) for samples dissolved in lysis buffers. 4. BSA (Bio-Rad) is used as a protein standard for both protein assays.

UV-spectral photometer e.g., Beckman DU-64 (see Note 2). 3. Methods 3.1. S. cerevisiae Cell Culture 1. The S. cerevisiae haploid wild type strain N318C is cultivated at 30 °C in defined synthetic YNB medium. First a preculture is grown, in which about 105 cells are inoculated in 30 mL culture medium (see previous section) and grown to a cell density of OD 0.6 measured at =610nm (see Note 3). 2. Then the main culture (30 mL, 300-mL Erlenmeyer flasks) is started with an inoculum from 10 μL out of the preculture. The cells are harvested by centrifugation (5,000g, 15 min) in the late midlogarithmic phase, when the culture reached an OD of 1.1 followed by washing in ice-cold sterilized water (see Note 4). 3. A control smear is performed on a Sabouraud agar plate to check for cellular contaminations. The incubation of the agar plate was done for 48 h at 37 °C in sterile conditions. 4. Harvested yeast cells are then transferred into individual 1.5 mL Eppendorf tubes and stored at –78 °C, if not processed immediately.

3.2. S-35 In Vitro Labeling (optional) 1. The S-35 in vitro labelling can be done additionally to the standard cell growth procedure. For the in vitro S 35 - labelling experiment the main culture is grown to an OD of 1.0 (=610 nm) 2. Then 10 mL of the cell suspension is inoculated with 100 μCi and kept shaking for a quarter of the fungal generation time (for S. cerevisiae in YNB – medium→ 30min). 3. After the S-35 incubation the cells are harvested by centrifugation (5 min, 5,000g) and the supernatant is discarded to the radioactive waste. The amount of protein, which will be extracted out of the obtained S-35 labeled pellet will be approx 5 mg protein solubilized in lysis buffer. Only methionine deficient mediums are suited for S-35 in vitro labeling.

3.3. Cell Disruption and Protein Solubilization 1. The yeast cell pellet from 10 mL culture OD 1.1 (see Section 21.3.1) is resuspended in 200 μL hot (95 °C) SDS buffer for presolubilization (see Note 5 and 6).

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2. After brief vortexing the hot suspension is sonicated 10 times for 1s (60 W, 20 kHz). The sonication has to be performed in such way that the probe is dipping as deep as possible in the suspension and not touching the sample cup during the agitation time. A 1.5-mL cup (conic bottom) is used for maximum abrasive agitation. Avoid foaming of the sample during this process (see Note 7). 3. After cell lyses the sample is boiled additionally for 5 min and then cooled down in an ice cold water bath to a sample temperature of maximum 20 °C (see Note 8 and 9). 4. Then the suspension is diluted with 500 μL lysis buffer and kept shaking for 20 min at room temperature. 5. After spinning down for 5 min at 10,000g, the protein concentration is measured from the clear supernatant (see Section 21.3.4). 6. The clear supernatant is stored at –78 °C or subjected to 2D-electrophoresis. An example of four replicate gels (IPG 4-7, 250 μg protein load, S. cerevisiae, SDS presolubilisation) is shown in Fig. 2 compared to a “standard preparation protocol” without the SDS presolubilization step (Fig. 1).

1

97kD

2

67kD 45kD

29kD

21kD

12kD 6kD

3

4

Fig. 1. Displaying four replicate 2D gels (IPG 4-7L, 250 μg protein load, fluorescence stain) from the yeast proteome (S. cerevisiae) performed without the SDS presolubilization step (see Note 12 and 13).

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5

Harder 97kD

6

67kD 45kD

29kD

21kD

12kD 6kD

7

8

Fig. 2. Displaying four replicate 2D gels (IPG 4-7L, 250 μg protein load, fluorescence stain) from the yeast proteome (S. cerevisiae) performed with the SDS - presolubilization step (see Note 14).

7. An approximate 7 μg/μL protein concentration can be expected in that obtained 700-μL sample volume (see Note 10)

3.4. Scintillation Counting and Protein Assay 1. 1. For S35 - labeled cells, the protein concentration is measured by removing 1 μL of the lysis buffer extract to a filter. After drying the sample, the filter is inserted into a counting vial, which is filled with 4.5 mL POPOP - solution. The scintillation counter is calibrated to a C12 – standard, which was first solubilized in lysis buffer. The scintillation counting should be repeated three times for each sample. Approximately 3 × 106 cpm should be loaded onto an IPG strip 4–7, 2.5 × 106 counts onto the gradient 3–10. This method is working with and without the SDS presolubilization step (see Note 11). 2. 20 μL of the SDS extract are used for the Lowry protein assay. 5 μL of the lysis buffer extract are taken for the Bradford protein assay.

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4. Notes 1. For S35 counting: after an appropriate amount of sample is pipeted onto the filter, the sample has to be dried for at least 30 min, due to water residues suppress the radio signal significantly. 2. The determination of the protein content is optional, but an optimum and reproducible protein load is essential for successful protein quantification. 3. Significantly increased proteolytic activity in fungi is observed in cells grown in minimal medium compared to cells grown in full medium, in cells harvested in their stationary phase (increases proteases activity to a level at least 100 times that of log phase cells), in vital cells stored or grown in nitrogen starvation and in the presence of peptone in the culture medium. 4. When you are researching fungal stress responses, washing the pellet with ice cold water will induce a deficient cell response. In that case the cell washing can be performed with 15 °C water as well. 5. The whole procedure is carried out in one single Eppendorf tube. By establishing the yeast sample preparation procedure main focus was given to its reproducibility, robustness, and maximum protein solubility with respect to the yeast’s peculiarities like proteolytic activity and cell wall constitution. 6. Dried pellets (for measuring the amount of cells) are not suited for further proteome analysis, due to heat shock responses of the cells. Homologue pellets should be used for protein extraction and 2D separation. 7. The correct sonication will provide a quantitative cell disruption and protein disaggregating by simultaneously cracking the fungal DNA and RNA strands, which otherwise can produce horizontal streaks in the 2D PAGE. The DNA and RNA fragments should become visible as a thin white foam after the sonication step. 8. By presolubilization in hot SDS buffer fungal proteases are denatured and inactivated during the cell breakage. Boiling the sample in SDS will increase protein solubility, especially for the high molecular weight and hydrophobic proteins, due to the high affinity of SDS to proteins in solution. 9. It is crucial to cool down the SDS-boiled sample below 20 °C, to avoid carbamylation reactions with the urea containing lysis buffer. The final SDS concentration should not exceed 0.25% in the extract to be applied onto the IPG strip, due to the interference of the SDS with the isoelectric focussing, so be sure that during the SDS boiling step the total volume is kept. 10. By starting with about 1 × 108 cells (S. cerevisiae) the obtained protein concentration will be approximately 7μg/μL in a 700 μL volume. 11. The protein concentration within this preparation method can be measured with the Lowry Kit after the SDS-presolubilization or with a scintillation counter (by S-35 labeled cells) out of the final extract. Determination of the dry weight or cell counting after cell harvesting should be done at least one time to check purity and approximate amount of cultivated cells.

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12. A major problem in fungal protein extraction is the strong fungal proteolytic activity. By applying a standard preparation protocol significant degradation can occur (see Fig. 1) and consequently a protein quantification is useless. 13. In many cases proteolytic activity is not as obvious as in the gels 1–4 (Fig. 1), especially when degradation only begins. It can show up in disappearance of single spots or in the decrease of the quantities of some spots, which can be spuriously interpreted as a cell response. Therefore the complete and irreversible inactivation of the mycotic proteome is essential. 14. Note, that the obtained “crude extract” will include proteins from the cytoplasm, the organelles, membrane bound proteins and surface proteins. Integral plasma membrane proteins as well as integral cell wall proteins are not extracted with that recipe (see Fig. 2).

References 1. Sougioultzis, S., Simeonidis, S., Bhaskar, K. R., et al. (2006) Saccharomyces boulardii produces a soluble anti-inflammatory factor that inhibits NF-kappaBmediated IL-8 gene expression. Biochem. Biophys Res Commun. 343, 69–76 2. Dalmasso, G., Loubat, A., Dahan, S., Calle, G., Rampal, P., and Czerucka, D. (2006) Saccharomyces boulardii prevents TNF-alpha-induced apoptosis in EHECinfected T84 cells. Res. Microbiol. 164, 876–84. 3. Hardwidge, P. R., Donohoe, S., Aebersold, R., and Finlay, B. B. (2006) Proteomic analysis of the binding partners to enteropathogenic Escherichia coli virulence proteins expressed in Saccharomyces cerevisiae. Proteomics. 6(7), 2174–79. 4. Jenkins, J. R., Pocklington, M. J., and Orr, E. (1990) The F1 ATP synthetase beta-subunit: a major yeast novobiocin binding protein. Cell Sci. Aug 96 ( Pt 4), 675–82. 5. Hermida, L., Brachat, S., Voegeli, S., Philippsen, P., and Primig, M. (2005) The Ashbya Genome Database (AGD) - a tool for the yeast community and genome biologists. Nucleic Acids Res. 33, 348–52. 6. [No authors listed]. (1997) The yeast genome directory.Nature 387 (6632 Suppl) 5. 7. Jones, E. W. (1991) Tackling the protease problem in Saccharomyces cerevisiae. Methods Enzymol. 19, 428–53. 8. Groll, M. and Huber, R. (2005) Purification, crystallization, and X-ray analysis of the yeast 20S proteasome. Methods Enzymol. 398, 329–36. 9. McIntyre, J., Podlaska, A., Skoneczna, A., Halas, A., and Sledziewska-Gojska, E. (2006) Analysis of the spontaneous mutator phenotype associated with 20S proteasome deficiency in S. cerevisiae. Mutat Res. 593(1–2), 153–63. 10. Garduno, E., Perez-Giraldo, C., Blanco, M.T., Hurtado, C., and Gomez-Garcia, A. C. (2005) Exposure to therapeutic concentrations of ritonavir, but not saquinavir, reduces secreted aspartyl proteinase of Candida parapsilosis. Chemotherapy 51(5), 252–5. 11. Schmidt, M., Haas, W., Crosas, B., et al. (2005) The HEAT repeat protein Blm10 regulates the yeast proteasome by capping the core particle. Nat. Struct. Mol. Biol. 4, 294–303.

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12. Mima, J., Hayashidam, M., Fujii, T., et al. (2005) Structure of the carboxypeptidase Y inhibitor IC in complex with the cognate proteinase reveals a novel mode of the proteinase-protein inhibitor interaction. J. Mol. Biol. 346(5), 1323–34. 13. Takai, T., Kato, T., Sakata, Y., et al. (2003) Recombinant Der p 1 and Der f 1 exhibit cysteine protease activity but no serine protease activity. Mol. Biol. 328(4), 944–52. 14. Fotedar, R. and Al-Hedaithy, S. S. (2005) Comparison of phospholipase and proteinase activity in Candida albicans and C. dubliniensis. Mycoses 48(1), 62–7. 15. Görg, A., Weiss, W., and Dunn, M. J. (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 04 (12), 3665–85. 16. Williams, B. and Wilson, K. (1994) Methoden der Biochemie. Thieme-Verlag Stuttgart. New York, 3. Auflage 1994 17. Johnson, T. M., Holady, s. K., Sun, Y., Subramaniam, P. S., Johnson H. M., and Krishna, N. R. (1999) Purification, and characterization of interferon-tau produced in Pichia pastoris grown in a minimal medium. Interferon Cytokine Res. 19(6), 631–6. 18. Harder, A., Wildgruber, R., Nawrocki, A., Fey, S. J., Larsen, P. M., and Gorg, A. (1999) Comparison of yeast cell protein solubilization procedures for twodimensional electrophoresis. Electrophoresis 20 (4–5), 826–9. 19. Wildgruber, R., Reil, G., Drews, O., Parlar, H., and Gorg, A. (2002) Web-based two-dimensional database of Saccharomyces cerevisiae proteins using immobilized pH gradients from pH 6 to pH 12 and matrix-assisted laser desorption/ionizationtime of flight mass spectrometry. Proteomics 2(6), 727–32. 20. Luche, S., Santoni, V., and Rabilloud, T. (2003) Evaluation of nonionic and zwitterionic detergents as membrane protein solubilizers in two-dimensional electrophoresis. Proteomics 3(3), 249–53. 21. Görg, A. Two-Dimensional Electrophoresis of Proteins using Immobilized pH Gradients: online manual at: http://www.weihenstephan.de/ blm/deg/manual/manfrm.htm 22. Rabilloud, T., Strub, J. M., Luche, S., Dorsselaer, A., and Lunardi, J. (2001) Comparison between Sypro Ruby and ruthenium II tris (bathophenanthroline disulfonate) as fluorescent stains for protein detection in gels. Proteomics 1(5), 699–704.

22 Isolation and Enrichment of Secreted Proteins from Filamentous Fungi Martha L. Medina and Wilson A. Francisco

Summary Filamentous fungi have been recognized as extraordinary producers of secreted proteins and are known to produce novel proteins and enzymes through dispensable metabolic pathways. Here, methods are described for the isolation and enrichment of samples of secreted proteins from cultures of filamentous fungi for analysis by gel electrophoresis and mass spectrometry techniques. These methods can be readily applied to the study of differential protein expression and secretion and metabolic pathways in filamentous fungi by proteomic approaches.

Key Words: Exoproteome; extracellular proteins; protein deglycosylation; protein precipitation; secreted proteins; secretome; gel electrophoresis.

1. Introduction Protein secretion plays an important role in filamentous fungi, particularly in nutrition, as secreted enzymes degrade complex biological molecules to serve as carbon and nitrogen sources. Filamentous fungi are known for their ability to secrete a broad spectrum of enzymes, the majority of which are hydrolytic, into the extracellular matrix (1). This ability has been widely exploited by the biotechnology industry for the production of enzymes for commercial and industrial use. Most commonly, filamentous fungi secrete proteins via a classical secretory pathway (2) and most, if not all, of the secreted proteins are glycosylated, containing modifications such as oligomannose N- and Oglycans (3). These attached sugars increase the stability of the secreted proteins From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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and provide resistance to environmental influences, as well as increase their solubility in the culture media. Although many studies of protein secretion have been carried out in yeast and animal systems, studies on protein secretion by filamentous fungi are limited (4). Typical studies have focused on the identification, purification and characterization of secreted proteins, but only a few studies have been conducted on the global analysis of fungal extracellular proteomes. Proteomic studies on the analysis of secreted proteins from a number of fungi, including Aspergillus flavus (5,6), A. oryzae (7), Fusarium graminearum (8), and Pleurotus sapidus (9), have recently appeared in the literature. Much of the delay in the use of proteomic techniques for the study of fungal protein expression in general can be attributed to the fact that there is a lack of complete and publicly available genome sequences from filamentous fungi. It is expected that as the number of published fungal genomes increases, proteomic studies on both intracellular and extracellular proteins will follow. Within their high capacity to produce secreted proteins, filamentous fungi are able to express and secrete proteins for “dispensable” metabolic functions (10). These enzymes participate in pathways that are either not required for growth or are only required for growth under a limited range of conditions. These changes in protein secretion provide an excellent platform for the systematic study of the overall protein secretion expression (secretome or exoproteome) as a function of culture conditions. Proteomic analysis using two-dimensional gel electrophoresis (2-DE) and mass spectrometry (MS) has proven to be the most powerful and sensitive method for the identification of proteins in complex mixtures. 2-DE provides an excellent platform to assess differential secreted protein expression. Although the number of proteins that can be analyzed by 2DE is still limited to 1,000–2,000 on one gel, the maximum number of secreted proteins by a filamentous fungi under a given set of conditions is well within this range, as a recent analysis of the genome of Aspergillus niger identified only about 400 putative secreted proteins from a total of about 5,100 genes (11). Obtaining MALDI-TOF MS data from tryptic digests of gel bands and spots and searching against online protein databases can identify 2-DE separated proteins. The success of peptide mass fingerprinting depends on the detection of a representative set of peptide masses derived from a protein and that the protein in question is known (it exists in a protein database). Alternatively, proteins can be identified by MS/MS with custom databases that contain sequences of all publicly known fungal proteins or by de-novo sequencing using this technique. Several technical issues need to be considered when studying the secretome of filamentous fungi by proteomic analysis. First, although fungi have the ability to secrete large amounts of protein, these proteins are highly diluted in the culture medium. Second, glycosylation poses a problem for visualization of proteins separated by gel electrophoresis, as glycosylated samples tend to

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smear in gels. In addition, glycosylation can protect proteins against proteolysis, making the identification of these proteins more difficult as peptide mass fingerprinting relies on efficient protease digestion before mass spectrometry (12). Also, the attached sugars greatly increase the size of generated peptide fragments, adding another layer of complexity in their identification by mass spectrometry. These problems can be overcome by concentrating the media, most commonly by lyophilization or ultrafiltration, and by removing the sugars with commercially available glycosidases or by chemical deglycosylation methods. Here we describe a general procedure for the isolation and enrichment of secreted proteins from cultures of filamentous fungi for proteomic analysis by gel electrophoresis and mass spectrometry, including protocols for protein deglycosylation. The methods described here can be easily adopted for the isolation and characterization of the secretome of filamentous fungi grown under varied conditions. 2. Materials 2.1. Filtration and Lyophilization 1. Miracloth (Calbiochem, EMD Biosciences, Inc., San Diego, CA) or No. 2 Whatman filter paper. 2. Lyophilizer

2.2. Centrifugation and Ultraﬁltration 1. Centricon Plus-70 Centrifugal Filter Device (Millipore, Billerica, MA). 2. Centrifuge

2.3. Precipitation with Trichloroacetic Acid 1. 20% (w/v) Trichloroacetic acid (TCA) (Sigma-Aldrich, St. Louis, MO), stored at 4°C. 2. 70% Ethanol, stored at –20°C. 3. Acetone

2.4. Precipitation with Methanol and Chloroform 1. Methanol 2. Chloroform

2.5. Enzymatic Deglycosylation 1. Ultrafree-0.5 centrifugal filter devices with Biomax-5 membranes 5,000 NMWL (Millipore, Billerica, MA).

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2. Peptide: N-glycosidase F (PNGase F) (New England BioLabs, Inc., Ipswich, MA). PNGase F is provided with 10× glycoprotein denaturing buffer (0.5% SDS, 1% -mercaptoethanol), 10× G7 reaction buffer (50 mM sodium phosphate, pH 7.5), 10% Nonidet P-40 (NP-40), and a solution of PNGase F (see Note 1).

2.6. Chemical Deglycosylation (see Note 2) 1. Reacti-Vial Reaction Vials (5 mL) (Pierce Chemical Co., Rockford, IL). 2. Trifluoromethanesulfonic acid (TFMS) (Sigma/Aldrich, St. Louis, MO, USA) (see Note 3). 3. Anisole, anhydrous (Sigma/Aldrich, St. Louis, MO), stored at 4°C. 4. 60% (v/v) Pyridine solution, stored at –20°C. 5. Diethyl ether, stored at –20°C. 6. 95% Ethanol, stored at 4°C.

2.7. SDS-PAGE 1. SDS Sample Reducing Buffer: Mix 4.05 mL of deionized water, 1.25 mL 0.5M Tris-HCl, pH 6.8, 2.50 mL glycerol, 2.00 mL 10% SDS and 0.20 mL 0.5% (w/v) bromophenol blue. 2. -Mercaptoethanol

2.8. Two-Dimensional Gel Electrophoresis 1. 2-DE Buffer: 8M urea, 2% (w/v) CHAPS, 50 mM dithiothreitol, 0.2% (w/v) 100× Bio-Lyte 3–10 ampholytes (Bio-Rad Laboratories, Inc., Hercules, CA), 0.001% (w/v) bromophenol blue.

3. Methods This chapter describes general-purpose sample preparation methods for the isolation and enrichment of secreted proteins from cultures of filamentous fungi. The methods described in the following section can be divided into (1) isolation and concentration by filtration and lyophilization or ultrafiltration, (2) precipitation, and (3) deglycosylation of concentrated protein samples by enzymatic (PNGase F) or chemical deglycosylation using TFMS acid (13). The samples can then be further analyzed by gel electrophoresis and mass spectrometry by established techniques described in other chapters. A schematic representation of these protocols is shown in Fig. 1. 3.1. Isolation and Concentration of Supernatant Broth 1. After the studied filamentous fungus has been grown in liquid culture media for the desired time, the broth containing the secreted proteins is collected by filtration

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Fig. 1. Simplified schematic for the isolation and enrichment of secreted proteins from cultures of filamentous fungi for proteomic analysis (see Methods section for details). through a Miracloth or No. 2 Whatman filter paper. Alternatively, the fungal mycelia can be separated from the supernatants by centrifugation at 10,000g for 10 min. (see Note 4). 2. The supernatants are concentrated by lyophilization (steps 3–5) or ultrafiltration (steps 6–8). 3. For lyophilization, place the filtered supernatants in individual round bottom flasks and freeze in liquid nitrogen. (see Note 5).

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4. Place the round bottom flasks on the lyophilizer until they are completely dry. 5. Redissolve the contents of the round bottom flasks in a minimal amount of deionized water, and store at –20°C until further analysis. 6. For ultrafiltration, place the filtered supernatant in a Centricon Plus-70 centrifugal filter device to a maximum of 70 mL. 7. Centrifuge at 3,500g until the desired final volume is achieved. 8. The concentrated supernatant can be washed with an appropriate buffer (e.g., 50 mM phosphate buffer, pH 6.0) and reconcentrated to remove excess salts, pigments, and metabolites. (see Note 6).

3.2. Protein Precipitation Protein precipitation is an efficient method for the removal of most contaminants, including detergents, salts, peptides, lipids, and phenolic compounds from protein samples. Either one of the methods described below can be used to precipitate the secreted proteins from the concentrated broth supernatant in preparation for gel electrophoresis analysis. It should be noted that no method will precipitate all proteins and some proteins will be difficult to resuspend following precipitation. 3.2.1. Precipitation with Trichloroacetic Acid (see Note 7) 1. Pipet 0.3 mL sample of concentrated broth supernatant into a 1.5-mL siliconized microcentrifuge tube and add an equal amount of cold TCA solution. 2. The mixture is incubated for 2 h at –20°C to allow the proteins to precipitate. After 2 h, allow samples to thaw if lightly frozen. 3. Centrifuge for 10 min at 14,000g. 4. The samples are decanted, and 1 mL of cold 70% ethanol is added, vortexed, and recentrifuged for 3 min. 5. Step 4 is repeated 3 times. 6. To completely dry the secreted protein pellet, add 1 mL of acetone, vortex and centrifuge for 1 min. The acetone is decanted, and the pellet is allowed to air dry for 30 min. 7. The dried pellet can be stored at –20°C until further analysis. 8. For electrophoresis analysis, the pellet is dissolved in SDS-PAGE or 2-DE sample buffer, as described below (see Subheading 3.5 and Note 8).

3.2.2. Precipitation with Chloroform/Methanol 1. To 100 μL of the concentrated supernatant in a siliconized microcentrifuge tube, add 400 μL of methanol, 100 μL of chloroform, and 300 μL of H2 O, and mix well. 2. Incubate at 4°C for 5 min and centrifuge at 9,000g at 4°C for 2 min.

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3. The upper phase is carefully removed and discarded. Add another 300 μL of methanol to the rest of the lower chloroform phase and the interphase with the precipitated protein and mix well. 4. Incubate at 4°C for 5 min and pellet the proteins by centrifugation at 13,000g for 5 min at 4°C. The supernatant is removed and the protein pellet is dried under a stream of air. 5. The dried pellet can be stored at –20°C until further analysis. 6. For electrophoresis analysis, the pellet is dissolved in SDS-PAGE or 2-DE sample buffer (see Subheading 3.5).

3.3. Protein Deglycosylation 3.3.1. Enzymatic Protein Deglycosylation with PNGase F 1. Concentrate a 500 μL sample of the concentrated supernatant broth to 50 μL using a Ultrafree-0.5 with a Biomax-5 membrane (5,000 NMWL) centrifugal filter device by centrifugation at 12,000g at 4°C. To desalt the sample, add 400 μL of deionized water to the concentrated sample and centrifuge again until a final volume of 50 μL is reached. 2. The concentrated, desalted solution is removed from the centrifugal filter device and placed in a 600 μL siliconized microcentrifuge tube. 3. Enzymatic digestion is carried out as follows and according to the manufacturer’s instructions (New England BioLabs, Inc.): 15 μL of denaturing buffer is added to the sample and boiled at 100°C for 10 min. To this sample, 3.5 μL of G7 buffer and 3.5 μL of NP-40 buffer are added, and the samples are digested with 1,000 U of PNGase F (see Note 9) for 18 hours at 37°C. 4. Following deglycosylation, the samples are precipitated as described above (see Subheading 3.3).

3.3.2. Chemical Protein Deglycosylation with Triﬂuoromethanesulfonic Acid 1. Glycoprotein samples should be relatively free of salts, minerals and detergents. Samples must also be completely dried. Lyophilize protein sample in a 5-mL Reacti-vial. 2. Incubate lyophilized sample with 0.3 mL of cold anisole and 0.6 mL of cold TFMS at 0°C in an ice-bath for 4 h under nitrogen with occasional shaking. 3. The reaction mixture is cooled to below –20°C by placing in a dry ice-ethanol bath and neutralized by slowly adding 1.2 mL of cold 60% aqueous pyridine solution. 4. The deglycosylated peptides are freed of reagents and low-molecular weight sugars by adding 2 mL of cold diethyl ether. The suspension is vortexed and extracted twice with cold ether.

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5. The aqueous layer is lyophilized, redissolved in deionized water, and precipitated as described in Subheading 3.3.

3.4. Preparation of Samples for Gel Electrophoresis Analysis (see Note 10) 3.4.1. SDS-PAGE 1. Add 50 μL of -mercaptoethanol to 950 μL SDS Reducing Buffer before use. 2. Dissolve the precipitated protein pellet in the appropriate volume of SDS reducing sample buffer determined by the size of the gel and the system used. 3. Boil sample for 4 min.

3.4.2. Two-dimensional Gel Electrophoresis 1. Dissolve the precipitated protein pellet in the appropriate volume of 2-DE Buffer determined by the size of the IPG strip and the system used for isoelectric focusing (see Note 11). 2. Incubate the sample for 2 h at room temperature and remove any insoluble material by centrifugation at 10,000g for 10 min.

4. Notes 1. PNGase F is an amidase that cleaves between the innermost N-acetylglucosamine and asparagine residues of high mannose, hybrid, and complex oligosaccharides from N-linked glycoproteins. PNGase F is the enzyme of choice for removing most N-linked oligosaccharides. Although secreted proteins could contain O-linked oligosaccharides, a recent study done on commercial cellulose enzyme preparation from the filamentous fungus Trichoderma reesei demonstrated that PNGase F treatment was superior to other enzymatic or chemical deglycosylation treatements in terms of yielding peptides through MALDI-MS that resulted in actual protein identification when searched against a database (12). 2. For chemical deglycosylation, the use of a commercially available kit is recommended. Two of such kits are the GlycoProfile IV, Chemical Deglycosylation Kit from Sigma/Aldrich (Sigma/Aldrich, St. Louis, MO, USA) and Glycofree Chemical Deglycosylation Kit from ProZyme, Inc. (San Leandro, CA, USA). 3. Trifluoromethanesulfonic acid is a strong acid, highly corrosive and hygroscopic. Protective goggles, laboratory coat and gloves should be worn when working with TFMS. 4. Typically, centrifuging is used to separate cells, in this case mycelia, from the supernatants. However, centrifuging does not work as well for filamentous fungi, as mycelia do not pack well and can float to the surface of the supernatant; therefore, filtering is preferred over centrifugation.

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5. Culture supernatants in the round bottom flasks should be frozen by spinning the flask while immersing in liquid nitrogen to ensure even distribution of the supernatant in the flask. This will allow better lyophilization of the samples. 6. An alternate method for desalting is dialysis. Concentrated supernatant broth can be dialyzed against water or any appropriate buffer at 4°C. 7. An alternate protocol for precipitation of fungal secreted proteins by trichloroacetic acid has been described by Suárez et al. (2005) (14). In this protocol, the concentrated, dialyzed and lyophilized culture filtrate is resuspended in 20% TCA in acetone containing 0.2% dithiothreitol (DTT), stored at –20°C. The suspension is kept at –20°C overnight. The sample is centrifuged at 16,000g at 4 C for 10 min and the resulting pellet is washed three times with acetone containing 0.2% DTT. The supernatants are removed by centrifugation and the pellet is dried overnight at room temperature. The pellet is dissolved in SDS-PAGE or 2-DE sample buffer before gel electrophoresis analysis (see Subheading 3.5). In this protocol, acetone is used to increase the solubility of interfering organic compounds and increase protein precipitation and DTT is included to prevent protein modification. 8. TCA precipitation allows for further concentration of the proteins in the sample, as well as for removal of non-protein substances, salts and other agents that may interfere with electrophoresis separation. Care should be taken to make sure the protein pellet is completely dry before adding sample buffer, as any leftover TCA will turn the Bromophenol Blue in the sample buffer yellow. 9. The majority of the proteins secreted by filamentous fungi can have carbohydrate contents reaching up to 50% of the total molecular weight of the protein (1). This heavy glycosylation is thought to be responsible for the tendency of these proteins to show “smearing” on SDS-PAGE gels (12). Enzymatic protein deglycosylation using PNGase F or chemical deglycosylation using TFMS acid will help to obtain better resolution of the protein bands on the SDS-PAGE and 2-DE gels. Deglycosylation can also aid in identification of proteins by MALDI-MS by obtaining better-resolved peaks in the mass spectra. 10. The preparation of samples for gel electrophoresis requires prior knowledge of protein concentration to determine the amount of protein to be loaded. The protein concentration in the concentrated supernatant broths can be determined according to the method of Bradford (15) using the Bio-Rad Protein Assay Reagent (Bio-Rad Laboratories, Inc., Hercules, CA), or any other commercially available kit. As fungal culture broths may contain phenolic compounds, it is important to note that many of these compounds interfere with several of the most popular methods for protein determination. It is suggested that protein concentration is determined following protein precipitation and resolubilization in SDS Reducing Buffer or 2-DE sample buffer. Unfortunately, several components of these buffers may also cause problems in the assessment of protein concentration. Two commercially available protein determination kits that can be used with samples prepared for electrophoresis techniques are 2-D Quant

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Kit (GE Healthcare Bio-Sciences Corp., Piscataway, NJ) and Advanced Protein Assay (Cytoskeleton, Inc., Denver, CO). 11. Resolubilization of the precipitated protein pellet in 2-DE buffer may require vortexing and/or sonication. It has been recently shown that adding 20–30 μL of 0.2M NaOH to the TCA precipitated pellet for 2 min, before adding the solubilization buffer, increases the amount of soluble protein in the sample buffer (16).

Acknowledgments The techniques described here were adapted from the appropriate original papers. This work was supported by NSF grant MCB-0317126 to W.A.F. M.L.M. was supported in part by a fellowship through the Research Training Group in Optical Biomolecular Devices provided under NSF grant DBI9602258-003 and the NSF-funded MGE@MSA Program, an Alliance for Graduate Education and the Professoriate, headquartered at Arizona State University. References 1. Peberdy, J. F. (1994) Protein secretion in filamentous fungi - trying to understand a highly productive black-box, Trends Biotechnol. 12, 50–7. 2. Conesa, A., Punt, P. J., van Luijk, N., and van den Hondel, C. A. A. J. J. (2001) The secretion pathway in filamentous fungi: A biotechnological view, Fungal. Genet. Biol. 33, 155–71. 3. Archer, D. B. and Peberdy, J. F. (1997) The molecular biology of secreted enzyme production by fungi, Crit. Rev. Biotechnol. 17, 273–306. 4. Wallis, G. L. F., Swift, R. J., Hemming, F. W., Trinci, A. P. J., and Peberdy, J. F. (1999) Glucoamylase overexpression and secretion in Aspergillus niger: analysis of glycosylation, BBA-Gen. Subjects 1472, 576–86. 5. Medina, M. L., Haynes, P. A., Breci, L., and Francisco, W. A. (2005) Analysis of secreted proteins from Aspergillus flavus, Proteomics 5, 3153–61. 6. Medina, M. L., Kiernan, U. A., and Francisco, W. A. (2004) Proteomic analysis of rutin-induced secreted proteins from Aspergillus flavus, Fungal. Genet. Biol. 41, 327–335. 7. Zhu, L. Y., Nguyen, C. H., Sato, T., and Takeuchi, M. (2004) Analysis of secreted proteins during conidial germination of Aspergillus oryzae RIB40, Biosci. Biotech. Bioch. 68, 2607–12. 8. Phalip, V., Delalande, F., Carapito, C., et al. (2005) Diversity of the exoproteome of Fusarium graminearum grown on plant cell wall, Curr. Genet. 48, 366–79. 9. Zorn, H., Peters, T., Nimtz, M., and Berger, R. G. (2005) The secretome of Pleurotus sapidus, Proteomics 5, 4832–38. 10. Keller, N. P. and Hohn, T. M. (1997) Metabolic pathway gene clusters in filamentous fungi, Fungal. Genet. Biol. 21, 17–29.

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11. Semova, N., Storms, R., John, T., et al. (2006) Generation, annotation, and analysis of an extensive Aspergillus niger EST collection, BMC Microbiol. 6:7. 12. Fryksdale, B. G., Jedrzejewski, P. T., Wong, D. L., Gaertner, A. L., and Miller, B. S. (2002) Impact of deglycosylation methods on two-dimensional gel electrophoresis and matrix assisted laser desorption/ionization-time of flight-mass spectrometry for proteomic analysis, Electrophoresis 23, 2184–93. 13. Edge, A. S. B., Faltynek, C. R., Hof, L., Reichert, L. E., and Weber, P. (1981) Deglycosylation of glycoproteins by trifluoromethanesulfonic acid, Anal. Biochem. 118, 131–7. 14. Suarez, M. B., Sanz, L., Chamorro, M. I., et al. (2005) Proteomic analysis of secreted proteins from Trichoderma harzianum - Identification of a fungal cell wall-induced aspartic protease, Fungal Genet. Biol. 42, 924–34. 15. Bradford, M. M. (1976) Rapid and sensitive method for quantitation of microgram quantities of protein utilizing principle of protein-dye binding, Anal. Biochem. 72, 248–54. 16. Nandakumar, M. P., Shen, J., Raman, B., and Marten, M. R. (2003) Solubilization of trichloroacetic acid (TCA) precipitated microbial proteins via NaOH for two-dimensional electrophoresis, J. Proteome Res. 2, 89–93.

23 Isolation and Solubilization of Cellular Membrane Proteins from Bacteria Kheir Zuobi-Hasona and L. Jeannine Brady

Summary Membrane proteins are rarely identified in two-dimensional electrophoretic (2-DE) proteomics maps. This is because of low abundance, poor solubility, and inherent hydrophobicity. In this study, membrane preparations from the Gram-positive bacterium Streptococcus mutans were isolated from protoplasts and by mechanical grinding. Membrane proteins were extracted using a mixture of trifluroethanol and chloroform, solubilized using highly chaotropic buffer containing ASB-14 and Triton X-100 and subjected to two-dimensional gel electrophoresis.

Key Words: Gram-positive; membrane proteins; solubilization; streptococcus mutans; two-dimensional gel electrophoresis.

1. Introduction Membrane proteins are notably limited in proteomic analysis of bacteria (1–4). The primary reason is their intrinsically hydrophobic nature leading to poor solubility (1,5). This hydrophobicity likely causes self-aggregation during isoelectric focusing leading to poor resolution and streaking in 2-D gels. Many approaches have been reported to improve recovery of hydrophobic proteins including fractionation for enrichment of proteins of interest and reduction of sample complexity (6,7), experimenting with different detergents to extract and solubilize these proteins (8–10), and extraction using organic solvents (11–13). While extraction of proteins from eukaryotic membranes and analysis by 2D electrophoresis has been technically feasible, such an approach to analyze From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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bacterial membrane proteins, particularly those from Gram-positive organisms has proved much more challenging. In this study membrane-associated proteins of the Gram-positive bacterium Streptococcus mutans were prepared using two techniques, one by mechanical grinding (14) and the other from protoplasts (15, 16). Both methods involve differential centrifugation to enrich for membranes within the preparation. The membrane associated proteins were extracted using a mixture of trifluroethanol and chloroform (13). This treatment produced three separate phases: An upper aqueous phase containing most of the proteins, an insoluble interphase containing few proteins and a lower chloroformic phase containing low concentration of proteins with low molecular weight (17). The extracted proteins were solubilized using highly chaotropic buffer containing ASB-14 and Triton X-100 and subjected to two-dimensional gel electrophoresis (Fig. 1). These techniques were efficient and highly reproducible for analysis of membrane proteins in S. mutans and suggest the potential use for other streptococci and Gram-positive bacteria in general.

2. Materials 2.1. Cell Culture and Lysis 1. 2. 3. 4.

UA-159, Streptococcus mutans, wild-type strain. Todd-Hewitt broth supplemented with 0.3% yeast extract (THYE). TM buffer: 50 mM maleate buffer, pH 6.0, containing 20 mM MgCl2 . Protease inhibitor cocktail (Sigma), reconstitute with DMSO and water, aliquot and store at –20°C. 5. Alumina Powder (A-5, Sigma). 6. Prechilled mortar and pestle (–20°C). 7. Wash buffer: 20 mM Tris-HCl, pH 6.8, containing 1 mM MgCl2 and protease inhibitor cocktail at a 1:200 v/v ratio.

2.2. Preparation of Membranes from Protoplasts 1. Phosphate-buffered saline (PBS). 2. Buffer A: 20% sucrose, 20 mM Tris-HCl, pH 7.0, and 10 mM MgCl2 . Store at 4°C. 2. Mutanolysin (INC Corp), store at -20°C. 3. Lysozyme (Sigma), store at –20°C. 4. Buffer B: 10 mM Tris-HCl, pH 8.1, 50 mM MgCl2 and 10 mM Glucose. Store at 4°C. 5. Buffer C: 10 mM Tris-HCl, pH 8.1, 50 mM NaCl and 20 mM MgCl2 . Store at room temperature. 6. Buffer D: 20 mM Tris-HCl, pH 7.2, and 10 mM MgCl2

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2.3. Triﬂuroethanol/Chloroform Extraction and Analysis 1. 2. 3. 4. 5.

50 mM ammonium bicarbonate pH 11.0, store at 4°C. Protease inhibitor cocktail (Sigma). Trifluroethanol/chloroform mixture (2:1 v/v). Ready Prep 2-D Cleanup Kit (Bio-Rad). Solubilization buffer: 7M urea, 2M thiourea, 2% Triton X-100, 0.5% ASB-14, 50 mM dithiothreitol (DTT) and 0.2% Bio-Lytes pH 3–10. Aliquot and store at –70°C. 6. RC DC protein assay kit (Bio-Rad). 7. Standard electrophoresis equipment from Bio-Rad: PROTEAN IEF Cell, Criterion Cell.

3. Methods 3.1. Cell Culture and Lysis 1. Transfer 100 mL of overnight culture of Streptococcus mutans, strain UA159 into 2 L of prewarmed Todd-Hewitt broth supplemented with 0.3% yeast extract. (THYE). 2. Incubate at 37°C with gentle agitation, until an absorbance reading of 0.7 at 600 nm is reached. 3. Place the culture immediately on ice for 20 min. 4. Harvest the cells by centrifugation at 12,000g for 10 min at 4°C. 5. Wash the cell pellet twice in TM buffer containing protease inhibitor at 1:30 v/v ratio. 6. Resuspend the cell pellet with 6 mL of the same buffer divide into three aliquots and store as slurries (see Note 1). 7. Transfer each frozen slurry to a prechilled mortar. 8. Add 3-g Alumina powder (A-5, Sigma). (see Note 2) 9. Use a prechilled pestle to grind the cells for 20 min. 10. Transfer the mixture into a 15 mL conical tube and centrifuge at 3,000g for 5 min at 4°C to remove the Alumina. 11. Centrifuge the supernatant at 16,000g for 10 min at 4°C to remove intact cells and cell debris. Discard the pellet. 12. Ultracentrifuge the supernatant at 100,000g for 18 h at 4°C to pellet the membranes. 13. Rinse the pellet three times with wash buffer (total volume 15 mL). 14. Resuspend the pellet in 5 ml wash buffer and centrifuge at 45,000g for 5 h at 4°C. 15. Remove the supernatant by pipette; it represents the cell cytoplasm and store at –70°C. 16. Keep the pellet in the centrifuge tube covered with parafilm, place in a box and store at –70°C until used. This represents the membrane-containing fraction.

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(A)

(B)

Fig. 1. Silver stained 2-D gel of Streptococcus mutans. (A) Membrane proteins obtained using the grinding method. (B) Cytoplasmic proteins. The 2-D gel profile of cytoplasmic proteins showed an entirely different distribution of protein spots as compared with the gel of extracted membrane proteins. Nanoelectrospray quadrupole time of flight (QTOF)-tandem mass spectrometry (MS/MS) analysis of several protein spots isolated from this gel (B) identified proteins predicted to be cytoplasmically localized, including translation elongation factor (spot #8), the 10 kDa chaperone GroES (spot #9), and ribosome recycling factor (spot #10). None of the protein spots identified

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3.2. Preparation of Membranes from Protoplasts 1. Transfer 100 ml of overnight culture of Streptococcus mutans, strain UA159 into 2 L of prewarmed Todd-Hewitt broth supplemented with 0.3% yeast extract (THYE). 2. Incubate at 37°C with gentle agitation, until an absorbance reading of 0.7 at 600 nm is reached. 3. Harvest the cells by centrifugation at 12,000g for 10 min at 4°C. 4. Wash the pellet with phosphate-buffered saline (PBS). 5. Re-suspend the cells with 50 mL Buffer A containing 150 μL protease inhibitor cocktail, 1 mg mutanolysin and 18 mg lysozyme. Swirl gently. 6. Incubate the mixture for additional 3 h at 37°C, swirling gently every 10 min. 7. Examine by light microscope under high power (see Note 3). 8. Collect the protoplast at 12,000g for 10 min at 4°C. Store supernatant at –20°C. 8. Wash the protoplasts with 30 mL Buffer B. 10. Add 100 μL protease inhibitor cocktail and passage twice using French Press under 10,000 psi. 11. Remove debris and unbroken protoplasts by centrifugation at 6,000g for 10 min. at 4°C. 12. Ultra-centrifuge the supernatant for 30 min. at 45,000g at 4°C. 13. Label the supernatant as cytoplasmic proteins and store at –70°C. 14. Resuspend the pellet in 3 mL Buffer C at room temperature. 15. Ultracentrifuge at 100,000g for 45 min at 4°C. 16. Decant the supernatant and wash the pellet twice with 2 mL Buffer D and decant the wash. 17. Store the pellet at –70°C until used.

3.3. Triﬂuoroethanol/Chloroform Extraction and Analysis 1. Re-suspend each membrane pellet (from Sections 3.1 and 3.2) in 150 μL of 50 mM ammonium bicarbonate (pH 11), containing 15 μL protease inhibitor cocktail and vortex well for one minute. 2. Add 1 mL of trifluroethanol/chloroform mixture (2:1 v/v) and vortex well for one minute. 3. Maintain the mixture on ice for 1 h and vortex every 5 min for 10 s each time.

Fig. 1. (Continued) from membrane extracts were observed in the 2-D gel of cytoplasmic proteins. On the other hand QTOF/MS/MS analysis of several spots isolated from gel (A) revealed known membrane and surface-associated proteins, including Enolase (spot #1), Biotin carboxyl carrier protein (spot #2), LemA-like protein (spot #3), Glucose 6-phosphate isomerase (spot #4), Phosphoglycerate kinase (spot #5), Glyeraldehyde 3-phosphate dehydrogenase (spot #6) and 50S ribosomal protein L7/L12 (spot #7).

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4. Centrifuge at 10,000g for 5 min. at 4°C to separate into three phases. 5. Carefully transfer the aqueous upper phase and the chloroformic lower phase into separate microcentifuge tubes. Keep the middle phase in the original microcentrifuge. 6. Dry using vacuum centrifuge for 2 h. 7. Solubilize each residue with 200 μL solubilization buffer, vortex for 1 min; let it set at RT then repeat vortexing twice. 8. Precipitate membrane proteins with the Bio-Rad clean-up kit (see Note 4). 9. Re-suspend the resulting pellets in solubilization buffer (from step 7), vortex well as in step 7. 10. Centrifuge at 13,000g for 5 min at room temperature (see Note 5). 11. Measure the protein concentration using RC DC protein assay and adjust to ∼250 μg/mL solubilization buffer from step 7 (see Note 6). 12. Analyze membrane fraction by standard two-dimensional electrophoresis protocols and silver stain the resulting 2-D gels.

4. Notes 1. Passage the cells through a syringe few times. The slurries can be placed in weighing boats wrapped, and stored frozen at –70°C until needed to be used. 2. Use mask to avoid inhaling the Alumina, it’s recommended to do the grinding in an ice pocket to avoid protein degradation. 3. More than 90% of cell wall digestion occurs after 3 h of incubation. Continuous gentle agitation can speed the process. 4. This step should be done using centrifuge at 4°C. Avoid over-drying the pellets, as it will be difficult to resuspend, resulting in loss of certain proteins. 5. It is necessary to clarify the protein sample from insoluble particles that might cause streaking in 2D gel. The supernatant can be used directly for isoelectricfocusing (IEF) in IPG strips. Store any remaining protein sample at –70°C for later analysis. 6. RC DC Protein Assay (Bio-Rad cat # 500-0121) or Plus One 2-D Kit (Amersham) can be used for protein quantitation. Dilute as necessary with solubilization buffer to yield the desired quantity of protein (50–100 μg, when using silver stain).

References 1. Santoni, V., Molloy, M. P. and Rabilloud, T. (2000) Membrane proteins and proteomics: un amour impossible? Electrophoresis 21, 1054–70. 2. Molloy, M. P., Herbert, B. R., Slade, M. B., Rabilloud, T., et al. (2000) Proteomics analysis of the Escherichia coli outer membrane. Eur. J. Biochem. 267, 2871–88. 3. Nouwens, A. S., Cordwell, S. J. Larsen, M. R., Molloy, M. P. et al. (2000) Complementing genomics with proteomics: the membrane subproteome of Pseudomonas aeruginosa PAO1. Electrophoresis, 21, 3797–3809.

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4. Wilkins, M. R., Gasteiger, E., Sanchez, J. –C., Bairoch, A. and Hochstrasser, D. F. (1998) Two-dimensional gel electrophoresis for proteome projects: the effect of protein hydrophobicity and copy number. Electrophoresis , 19, 1501–05. 5. Pasquali, C. Fialka, I., and Huber, L. A. (1997) Preparative two-dimensional gel electrophoresis of membrane proteins. Electrophoresis 18, 2573–81. 6. Molloy, M. P., Herbert, B. R., Walsh, B. J., Tyler, M. I., et al. (1998) Extraction of membrane proteins by differential solubilization for separation using twodimensional gel electrophoresis. Electrophoresis 19, 2573–81. 7. Lehner, I., Niehot, M., and Borlak, J., (2003) An optimized method for the isolation and identification of membrane proteins. Electrophoresis 24, 1795–1808. 8. Rabilloud, T., Blisnick, T., Heller, M., Luche, S., et al. (1999) Analysis of membrane proteins by two-dimensional electrophoresis: comparison of the proteins extracted from normal or Plasmodium faciparum-infected erythrocyte ghosts. Electrophoresis 20, 3603–10. 9. Wissing, J., Heim, S., Flohe, L., Bilitewski, U., and Frank, R. (2000) Enrichment of hydrophobic proteins via Triton X-114 phase partitioning and hydroxyapatite column chromatography for mass spectrometry. Electrophoresis 21, 2589–93. 10. Wasinger, V. C., Pollack, J. B., and Humphery-Smith, I., (2000) The proteome of Mycoplasma genitalium. Chaps-soluble component. Eur. J. Biochem. 267, 1571–82. 11. Molloy, M. P., Herbert, B., Williams, K. L., and Gooley, A. A., (1999) Extraction of Escherichia coli proteins with organic solvents before two-dimensional electrophoresis. Electrophoresis 20, 701–4. 12. Seigneurin-Berny, D., Rolland, N., Garin, J., and Joyard, J., (1999) Technical Advance: Differential extraction of hydrophobic proteins from chloroplast envelope membranes: a subcellular specific proteomic approach to identify rare intrinsic membrane proteins. Plant. J. 19, 217–228. 13. Deshusses, J. M. P., Burgess, J. A., Scherl, A., Wenger, Y., Walter, N., et al. (2003) Exploitation of specific properties of trifluroethanol for extraction and separation of membrane proteins. Proteomics, 3, 1418–24. 14. Vadeboncoeur, C., St Martin, S., Brochu, D., and Hamilton, I. R., (1991) Effect of growth rate and pH on intracellular levels and activities of the components of the phosphoenolpyruvate: sugar phosphotransferase system in streptococcus mutans Ingbritt. Infect. Immun. 59, 900–6. 15. Wessel, D. and Flugge, U. I., (1984) A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal. Biochem. 138, 141–3. 16. Bordier, C. (1981) Phase separation of integral membrane proteins in Triton X-114 solution. J. Biol. Chem. 256, 1604–7. 17. Zuobi-Hasona, K., Crowley, P. J., Hasona, A., Bleiweis, A. S., and Brady, L. J. (2005) Solubilization of cell membrane proteins from Streptococcus mutans for two-dimensional gel electrophoresis. Electrophoresis 26, 1200–6.

24 Isolation and Solubilization of Gram-Positive Bacterial Cell Wall-Associated Proteins Jason N. Cole, Steven P. Djordjevic, and Mark J. Walker

Summary This chapter describes a simple, rapid and reproducible method to prepare bacterial cell wall extracts for two-dimensional gel electrophoresis (2DE). The extraction process uses mutanolysin, an N-acetylmuramidase, to gently solubilize cell wall-associated proteins from Gram-positive prokaryotes. The cells are first washed with buffer and resuspended in a solution containing mutanolysin. Following incubation at 37 °C, the sample is centrifuged and the supernatant containing the soluble cell wall-associated proteins is harvested. Following a brief precipitation step, the pellet is solubilized in sample buffer ready for isoelectric focusing and 2DE analysis.

Key Words: Bacterial proteome; cell wall-associated; Gram-positive; group A streptococcus; mutanolysin; Streptococcus pyogenes; two-dimensional gel electrophoresis.

1. Introduction Gram-positive bacteria are bounded by a thick cell wall composed of primarily of peptidoglycan, a large macromolecule of acetamido sugars and amino acids (1). The glycan chains of peptidoglycan consist of alternating units of N-acetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) in -1,4 linkage (2). Short cell wall peptides are cross-linked to the glycan chains to form a three-dimensional molecular network. The major function of the cell wall envelope is to provide a rigid exoskeleton for protection against mechanical and osmotic lysis. The cell wall also maintains a defined cell shape and plays an integral role in the anchoring of proteins to the cell surface (3,4). From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractionation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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Most Gram-positive cell walls are resistant to dissolution with lysozyme (5). Mutanolysin, a muralytic enzyme derived from Streptomyces globisporus 1829, cleaves the -1,4 linkage of the N-acetylmuramyl-N-acetylglucosamine in the glycan backbone of the peptidoglycan-polysaccharide polymer, which is highly conserved among bacterial species (6). Although mutanolysin has the same specificity as lysozyme, it has a wider range of activity among peptidoglycanpolysaccharide from bacterial strains resistant to lysozyme, including Streptococcus pyogenes (group A streptococcus), a Gram-positive pathogen responsible for numerous human diseases. In this chapter, the authors describe a convenient and efficient mutanolysin extraction method for the solubilization of cell wall-associated proteins from S. pyogenes. A detailed discussion is presented on bacterial growth conditions, cell wall purification, preparative SDS-PAGE analysis, sample preparation, isoelectric focusing and 2DE.

2. Materials Unless stated otherwise, all solutions were prepared with glass distilled water and high purity electrophoresis grade reagents. Review the manufacturers’ safety data sheets (MSDS) and follow the safety precautions and general handling procedures for hazardous materials. 2.1. Bacterial Culture 1. Horse blood agar plates (BioMérieux). Store at 4 °C. 2. THBY medium: 30 g/L Todd-Hewitt broth, 10 g/L yeast extract (Difco). Autoclave and store at room temperature.

2.2. Extraction of Cell Wall-Associated Proteins 1. TE buffer: 50 mM Tris-HCl, 1 mM EDTA, pH 8.0. Autoclave and store at 4 °C. 2. TE-Sucrose (TES) buffer: 20% (w/v) sucrose in TE buffer (pH 8.0). Autoclave and store at 4 °C. 3. Phenylmethylsulfonyl fluoride (PMSF, Sigma) dissolved at 1 mM in chilled TES buffer (see Note 1). PMSF is very toxic and rapidly degrades in aqueous solutions. Prepare immediately before use and place on ice. 4. Mutanolysin: Resuspend 10,000 units of chromatographically purified mutanolysin from Streptomyces globisporus ATCC 21553 (Sigma) in 2 mL of chilled filtersterilized 0.1 M K2 HPO4 (pH 6.2) for a working solution of 5,000 units/mL. Store 200 μL aliquots at –20 °C. 5. Lysozyme dissolved in chilled TES buffer at 100 mg/mL. Avoid foaming by gently pipeting up and down. Prepare fresh each time and store on ice before use.

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6. Mutanolysin mix: 1 mL TES buffer, 100 μL lysozyme (100 mg/mL in TES), 50 μL mutanolysin (5,000 U/mL in 0.1 M K2 HPO4 , pH 6.2). Prepare immediately before use and place on ice.

2.3. Bicinchoninic Acid (BCA) Protein Assay 1. Bicinchoninic Acid Protein Assay Kit (Sigma): Store Reagents A and B at room temperature. 2. Bovine serum albumin (BSA, Sigma). 3. TES buffer: 20% (w/v) sucrose in TE buffer (50 mM Tris-HCl, 1 mM EDTA, pH 8.0). Autoclave and store at 4 °C. 4. Microtiter plate reader capable of measuring absorbance in the 560 nm region. 5. Microtiter plate and sealing film.

2.4. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. 0.5 M Tris-HCl (pH 6.8) and 1.5 M Tris-HCl (pH 8.8) prepared in Milli-Q® water. Adjust pH with concentrated HCl. Store at 4 °C. 2. 10% (w/v) SDS dissolved in Milli-Q® water. Store at room temperature. 3. 40% acrylamide/bis solution (37.5:1, Amresco). Acrylamide monomer is a neurotoxin and suspected carcinogen; care should be taken to avoid exposure. Store in dark at 4 °C. 4. N,N,N ,N -Tetramethylethylenediamine (TEMED, Amresco). Store at room temperature. 5. Ammonium persulfate (APS, Amresco) prepared at 10% (w/v) in Milli-Q® water (see Note 2). 6. Gel overlay solution: 70% (v/v) ethanol. Store at room temperature. 7. 10× running buffer: 250 mM Tris-HCl, 1.92 M glycine, 1% (w/v) SDS, pH 8.3. Do not adjust pH with acid or base. Dilute to 1× working solution and mix thoroughly before use. Store at room temperature. 8. 1 M Dithiothreitol (DTT) dissolved in 0.01 M sodium acetate (pH 5.2). Filtersterilize and store 1 mL aliquots at –20 °C. 9. 5× loading buffer: 225 mM Tris-HCl (pH 6.8), 50% (v/v) glycerol, 5% (w/v) SDS, 0.05% (w/v) Bromophenol Blue, 250 mM DTT (see Note 3). Store at room temperature. Dilute sample 1:5 with loading buffer and heat at 95 °C for 10 min. 10. Molecular weight markers: PageRuler™ Protein Ladder (Fermentas). Store at –20 °C. 11. Coomassie™ Blue staining solution: 0.2% (w/v) Coomassie™ Blue R250, 40% (v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 12. Rapid destain solution: 40% (v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 13. Final destain solution: 4% (v/v) glycerol, 10% (v/v) acetic acid. Store at room temperature.

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2.5. Trichloroacetic Acid (TCA) Precipitation 1. 10% (v/v) TCA (Sigma): Protect from light and store at room temperature. TCA is a highly corrosive acid. Use safety equipment when handling to avoid exposure. 2. Ethanol 100% (v/v) analytical reagent stored at –20 °C. 3. Sample solubilization solution: 8 M urea, 100 mM DTT, 4% (w/v) CHAPS, 0.8% (v/v) Bio-Lyte® 3/10 ampholyte (Bio-Rad), 40 mM Tris-HCl. Make up to volume with Milli-Q® water (see Note 4). Store 1 mL aliquots at –20 °C. Thaw required number of aliquots and discard leftover solution.

2.6. Two-Dimensional (2D) Gel Electrophoresis 2.6.1. Sample Preparation and Rehydration 1. Sample solubilization solution: 8 M urea, 100 mM DTT, 4% (w/v) CHAPS, 0.8% (v/v) Bio-Lyte® 3/10 ampholyte (Bio-Rad), 40 mM Tris-HCl. Make up to volume with Milli-Q® water. Store 1 mL aliquots at –20 °C. Thaw required number of aliquots and discard leftover solution. 2. Water-bath sonicator or equivalent. 3. Bromophenol Blue solution 1% (w/v) in Milli-Q® water. Pass through a 0.22-μm filter unit (Millipore) to remove particulate matter and store at room temperature. 4. 11-cm ReadyStrip™ linear pH 4-7 IPG strips (Bio-Rad). 5. Disposable plastic rehydration tray (Bio-Rad). 6. Mineral oil (Bio-Rad). Store at room temperature.

2.6.2. First Dimension Isoelectric Focusing (IEF) 1. 2. 3. 4.

Electrode wicks (Bio-Rad). PROTEAN® IEF Cell and 11-cm focusing tray (Bio-Rad). Rehydrated 11-cm ReadyStrip™ linear pH 4–7 IPG strip (Bio-Rad). Mineral oil (Bio-Rad). Store at room temperature.

2.6.3. Casting 2D Gels 1. 1.5 M Tris-HCl (pH 8.8) prepared in Milli-Q® water. Adjust pH with concentrated HCl. Store at 4 °C. 2. 10% (w/v) SDS dissolved in Milli-Q® water. Store at room temperature. 3. 40% acrylamide/bis solution (37.5:1, Amresco). Store in dark at 4 °C. 4. TEMED (Amresco). Store at room temperature. 5. 10% (w/v) APS in Milli-Q® . Prepare fresh each time. 6. Water-saturated n-butanol gel overlay solution: Combine equal volumes of n-butanol and Milli-Q® water and shake to mix. Allow to settle and use top (aqueous layer) to overlay gels. Store at room temperature. 7. Gel storage solution: 0.375 M Tris-HCl (pH 8.8), 0.1% (w/v) SDS. Store at 4 °C.

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2.6.4. Second Dimension SDS-PAGE 1. Disposable plastic equilibration tray (Bio-Rad). 2. Equilibration buffer: 6 M urea, 2% (w/v) SDS, 375 mM Tris-HCl (pH 8.8), 20% (v/v) glycerol, 2.5% (v/v) acrylamide, 130 mM DTT. Store 4 mL aliquots at –20 °C. Thaw required number of aliquots and discard leftover solution. 3. Molecular weight markers: PageRuler™ Prestained Protein Ladder (Fermentas). Store at –20 °C. 4. 10× running buffer: 250 mM Tris-HCl, 1.92 M glycine, 1% (w/v) SDS, pH 8.3. Do not adjust pH. If the pH is not accurate remake buffer. Dilute to 2× and 1× working concentrations and mix thoroughly before use. Store at room temperature. 5. Bromophenol Blue solution 1% (w/v) in Milli-Q® water. Pass through a 0.22 μm filter unit (Millipore) to remove particulate matter and store at room temperature. 6. Agarose gel overlay solution: 1% (w/v) agarose in 1× running buffer (pH 8.3). Do not adjust pH. Add 0.1 mL of 1% (w/v) Bromophenol Blue per 100 mL of overlay solution and heat in a microwave oven until completely melted (see Note 5). Allow to cool slightly (60 °C) before using. Store at room temperature after use.

2.6.5. Protein Detection and Gel Documentation 1. Colloidal Coomassie™ stain: 17% (w/v) ammonium sulfate, 3% (v/v) phosphoric acid, 34% (v/v) methanol, 0.1% (w/v) Coomassie™ G250 (see Note 6). Store at room temperature. 2. Destain solution: 1% (v/v) glacial acetic acid. Store at room temperature.

3. Methods For full information pertaining to chemical and electrical hazards refer to the manufacturers’ material safety data sheets (MSDS) and instruction manuals. S. pyogenes (UN2814) is a potential human pathogen and safety precautions must be taken to avoid exposure. 3.1. Bacterial Culture 1. Sixteen-streak the S. pyogenes strain onto a fresh horse blood agar plate (see Note 7). Seal with Parafilm® and incubate at 37 °C overnight (approx 16 h). 2. Inoculate a single, well-isolated hemolytic colony (see Note 8) into a 5 mL sterile tube containing 2 mL of THBY medium. Incubate at 37 °C overnight for 16 h without shaking. 3. To a 250 mL conical flask containing 98 mL of THBY, add the entire volume of overnight culture and incubate at 37 °C without agitation until late stationary phase is reached (approx 16 h) (see Note 9).

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3.2. Extraction of Cell Wall-Associated Proteins 1. Transfer the overnight culture to a sterile centrifuge tube and harvest the bacterial cells by centrifugation at 7,560g for 20 min at 4 °C. 2. Carefully decant the culture supernatant and discard. Place the bacterial pellet on ice for 5 min. 3. Resuspend the pellet in 5 mL of chilled TE buffer containing 1 mM PMSF (see Note 10) by pipetting up and down. Take care to avoid foaming and ensure no bacterial clumps are visible. Centrifuge at 7,560g for 20 min at 4 °C and discard the supernatant. 4. Repeat step 3. Resuspend the pellet in 1.15 mL of ice-cold mutanolysin mix by pipeting up and down. Take care to avoid foaming and ensure no bacterial clumps are visible. 5. Transfer to a sterile microcentrifuge tube and incubate for 2 h at 37 °C with shaking (200 rpm) (see Note 11). 6. Centrifuge at 14,000g for 5 min at room temperature in a bench-top microcentrifuge. 7. Collect the supernatant (solubilized cell wall-associated proteins) by aspiration. Store 500 μL aliquots at –20 °C (see Note 12). 8. Determine the protein concentration of the cell wall extract using the BCA protein assay described below. 9. Prepare for SDS-PAGE analysis by diluting 20 μL of 5× loading buffer with 80 μL of cell wall extract in a microcentrifuge tube. Pierce the lid and boil for 10 min. Allow to cool to room temperature before loading (see Note 13).

3.3. BCA Protein Assay 1. Prepare the required amount of BCA Working Reagent by mixing 50 parts of Reagent A (Sigma proprietary solution containing BCA, sodium carbonate, sodium tartrate and sodium bicarbonate in 0.1 M NaOH, pH 11.25) with 1 part of Reagent B, a 4% (w/v) solution of copper(II) sulfate pentahydrate. Mix by vortexing until the solution is a uniform light green color (see Note 14). 2. Prepare BSA protein standards ranging from 0.2 to 1 mg/mL in TES buffer (see Note 15). Include a blank containing buffer with no protein. Protein standards may be stored at –20 °C. 3. Dilute the cell wall extract 1:5 in TES buffer to ensure the concentration is within the linear range of 0.2 to 1 mg/mL. 4. Add 25 μL of each standard, cell wall extract and blank to a microtiter plate before adding 200 μL of Working Reagent (see Note 16). 5. Seal the plate, incubate at 37 °C for 30 min and cool to room temperature (see Note 17). 6. Measure the absorbance at 562 nm with a microtiter plate reader and estimate the protein concentration of unknown samples from the BSA standard curve (see Note 18).

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3.4. SDS-PAGE This procedure is for use of the discontinuous Laemmli system (7) in a Mini-PROTEAN® 3 Cell (Bio-Rad) and can be readily adapted for other gel formats. 1. Thoroughly clean the glass plates with a laboratory detergent, rinse completely with distilled water and air dry. 2. Prepare a 12% resolving gel by mixing 3 mL of 40% acrylamide/bis solution with 2.5 mL of 1.5 M Tris-HCl (pH 8.8), 4.35 mL Milli-Q® water, 100 μL 10% (w/v) SDS, 15 μL TEMED and 50 μL 10% (w/v) APS (see Note 19). Pour a 0.75 mm thick resolving gel to the required level and immediately overlay with 70% (v/v) ethanol to give a flat gel surface (see Note 20). Allow the gel to polymerize for 45 min to 1 h at room temperature (see Note 21). The resolving gel can be submerged in 1.5 M Tris-HCl (pH 8.8) diluted 1:4 in distilled water and stored at 4 °C for up to 1 wk. 3. Decant the ethanol and thoroughly rinse the gel surface with distilled water. Remove excess water between the glass plates above the resolving gel with a piece of filter paper, taking care to avoid the gel surface. 4. Prepare the 4% stacking gel by mixing 0.5 mL of 40% acrylamide/bis solution with 2.5 mL of 0.5 M Tris-HCl (pH 6.8), 3.18 mL Milli-Q® water, 50 μL 10 % (w/v) SDS, 5 μL TEMED and 25 μL 10% (w/v) APS. Pour the stacking gel, insert the comb and allow the stacking gel to polymerize for 30–45 min at room temperature. 5. Prepare the running buffer by diluting 100 mL of 10× running buffer with 900 mL of distilled water. Mix thoroughly before use. 6. After the stacking gel has set, gently remove the comb and thoroughly rinse the wells with running buffer. Place the gel into the electrophoresis module and add running buffer to the upper and lower chambers. Load 15 μL of sample (approx 20 μg protein) and molecular weight markers for size comparison (see Note 22). 7. Connect the lid of the electrophoresis unit to a suitable power supply and run at a constant 200 V for approximately 45 min, or until the dye front reaches the bottom of the gel (see Note 23). 8. Once electrophoresis is complete, discard the running buffer and remove the stacking gel from the resolving gel. Place the resolving gel in a plastic tray and submerge in Coomassie™ Blue staining solution (see Note 24). Cover the tray to minimize exposure to vapors, microwave (600 W for 15 s) and shake at room temperature for 1 h or leave overnight. Decant stain (see Note 25) and add enough destain solution to completely cover the gel. Microwave for 15 s and shake for 20 min at room temperature. Repeat with fresh destain solution until the background is clear (see Note 26). 9. Discard destain and submerge gel in final destain solution ready for documentation with the GS-800™ calibrated densitometer (Bio-Rad) or equivalent (see Note 27). A representative SDS-PAGE gel of cell wall extracts from S. pyogenes is shown in Fig. 1.

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Fig. 1. Coomassie™ Blue stained 12% SDS-PAGE reducing gel of a mutanolysin cell wall extract harvested from S. pyogenes strain 5448 (serotype M1) after growth at 37 °C to late stationary phase in THBY medium without agitation. Molecular mass markers are given in kilo-Daltons (kDa).

3.5. TCA Precipitation 1. Add 1 mL of 10% (v/v) TCA to 1 mL of cell wall extract and mix immediately by vortexing at maximum output (see Note 28). Incubate on ice for 20 min. 2. Centrifuge at 14,000g for 15 min at room temperature in a bench-top microcentrifuge. 3. Discard supernatant and wash pellet with 1 mL of ice-cold 100% (v/v) ethanol to remove residual TCA (see Note 29). 4. Centrifuge at 14,000g for 5 min at room temperature in a bench-top microcentrifuge. 5. Discard the supernatant and dry the pellet for 30–60 min at room temperature. Store TCA precipitated pellets at –20 °C before 2DE (see Note 30).

3.6. 2D Gel Electrophoresis 3.6.1. Sample Preparation and Rehydration 1. Completely resuspend the TCA precipitated pellet in 500 μL of sample solubilization solution by pipeting up and down, taking care to avoid foaming (see Note 31).

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2. Sonicate at 14 W for 30 sec and vortex for 15 sec at room temperature to aid solubilization. Repeat this step a further 3 times (see Note 32). 3. Centrifuge at 14,000g for 15 min at room temperature in a bench-top microcentrifuge (see Note 33). 4. Transfer the supernatant to a new microcentrifuge tube and discard the pellet (see Note 34). 5. Calculate the volume of sample required for 200 μg protein and dispense into a new microcentrifuge tube. Adjust the volume to 250 μL with sample solubilization solution (see Note 35). 6. Add 2 μL of Bromophenol Blue solution and mix by briefly vortexing (see Note 36). 7. Pipette 250 μL of sample along the length of a channel in an 11 cm rehydration tray. Eliminate air bubbles using forceps to ensure even distribution of the sample in the IPG strip. 8. Carefully place the IPG strip onto the sample gel side down. Eliminate air bubbles under the strip by gently lifting the strip from one end with forceps. 9. Carefully overlay the strip with 2 mL of mineral oil to prevent evaporation (see Note 37). Attach the rehydration tray lid and rehydrate overnight (11–16 h) at room temperature (see Note 38). The sample will be passively absorbed by the strip during rehydration. 10. Remove the IPG strip from the rehydration tray with forceps (see Note 39). Remove the excess oil by holding the strip vertically and gently touching the tip of the backing strip to blotting paper (see Note 40).

3.6.2. First Dimension IEF The following IEF method assumes the use of the PROTEAN® IEF Cell (Bio-Rad) (see Note 41). 1. Equilibrate four electrode wicks per IPG strip in Milli-Q® water (see Note 42). 2. Remove excess water by briefly blotting the wicks on filter paper. Place two hydrated wicks directly on top of the cathode and anode electrode wires of the focusing tray. 3. Place the IPG strip gel side down onto the wicks, with the acidic (positive) end of the strip at the anode (positive) electrode of the focusing tray (see Note 43). 4. Overlay the IPG strip with 2 mL of mineral oil and eliminate air bubbles under the strip by gently lifting it from one end with forceps (see Note 44). Place the lid on the focusing tray and place into the IEF Cell, ensuring good contact between the electrodes of the focusing tray and the electrodes of the IEF Cell. 5. Program the IEF Cell with the following focusing conditions: 100 V for 1 h; 300 V for 1 h; 600 V for 1 h; 1,000 V for 1 h; 2,000 V for 1 h; 4,000 V for 40,000 Volt-hours (Vh) (approx 10 h); and 100 V hold (12 h) (see Note 45). Use an IEF Cell temperature of 20 °C with a maximum current of 50 μA per IPG strip.

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6. Remove the IPG strip from the IEF tray and drain the excess mineral oil with blotting paper as described in step 10 above. Proceed directly to the equilibration step (see Note 46).

3.6.3. Casting 2D Gels This protocol is for the casting of homogenous polyacrylamide gels with the Ettan™ DALTsix Gel Caster (GE Biosciences). 1. Lay the Ettan™ DALTsix Gel Caster on its back. Remove the faceplate and place the triangular rubber wedge in the V-shaped base. 2. Place a separator sheet against the back wall of the caster (square corners against the rubber wedge) followed by a 1.0 mm gel casting cassette. Fill the caster by alternately layering separator sheets and gel cassettes, ending with a separator sheet. Add thicker filler sheets until the stack of cassettes is flush with the edge of the caster (see Note 47). 3. Carefully place the faceplate onto the caster, clamp with six spring clips and tighten the faceplate screws (see Note 48). 4. Attach the cap to the filler port on the faceplate to prevent leakage. Place the Gel Caster in an upright position on a level surface ready for casting. 5. Prepare the 12.5% gel solution by mixing 156 mL of 40% acrylamide/bis with 125 mL of 1.5 M Tris-HCl (pH 8.8), 6 mL 10% (w/v) SDS and 208 mL Milli-Q® water. Mix well with a magnetic stirrer before adding 6 mL 10% (w/v) APS and 100 μL TEMED (see Note 49). To cast a homogenous 1.0-mm-thick gel, add the APS followed by TEMED and slowly pour the gel solution into the filling channel located at the back of the caster. Continuing pouring until the level is approx 4 cm below the top edge of the short plate (see Note 50). 6. Immediately overlay each gel with 4 mL of water-saturated n-butanol to exclude air and ensure a level surface. Cover the top of the Gel Caster with plastic wrap and allow gels to polymerize for 3–4 h at room temperature. 7. Pour off the gel overlay solution and rinse the surface of each gel with Milli-Q® water to remove residual n-butanol and unpolymerized acrylamide. Add 5 mL of gel storage solution to the top of each gel. Seal the top of the Caster with plastic wrap and store at 4 °C overnight (see Note 51).

3.6.4. Second Dimension SDS-PAGE The following procedure is for the running of large format (26 × 20 cm) acrylamide gels with the vertical Ettan™ DALTsix Electrophoresis Unit (GE Biosciences) (see Note 52). 1. Turn on the MultiTemp™ III heat exchanger and equilibrate to 10 °C. 2. Place the IPG strip gel side up into an 11 cm equilibration tray and add 2 mL of Equilibration buffer. Rock gently for 20 min at room temperature (see Note 53).

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3. During the equilibration step, rinse the glass cassettes with water to remove adhering acrylamide. Rinse the surface of each gel with Milli-Q® water and invert to drain (see Note 54). 4. Remove the IPG strips from the equilibration tray and wash each side with 3 mL of 1× running buffer (see Note 55). 5. Hold the gel cassette in a horizontal position with the short glass plate side up. Using forceps, place the IPG strip onto the long glass plate with the plastic backing against the plate. By carefully pushing against the plastic backing of

Fig. 2. Examination of the cell wall proteome of S. pyogenes strain 5448 (serotype M1). The mutanolysin cell wall extract was harvested after growth at 37 °C to late stationary phase in THBY medium without agitation. The extracts were concentrated by TCA precipitation, isoelectric focused over a pH range of 4–7 and resolved with a 12.5% SDS-PAGE gel. The gel was stained with colloidal Coomassie™ and several landmark proteins identified by MALDI-TOF peptide mass fingerprinting analysis. Identified protein spots are denoted by numbered arrows, which correspond to the proteins in Table 1. Molecular mass markers are given in kilo-Daltons (kDa).

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7. 8.

9.

10.

Cole et al. the strip with a spatula, slide the strip between the glass plates down to the gel surface. Take care not to damage the gel matrix with the spatula during this step. The strip should rest gently against the gel surface with the positive (acidic) end against the left edge of the glass plate (see Note 56). Apply 10 μL of prestained molecular weight markers to a piece of filter paper or electrode wick paper. Allow the markers to soak in and position on the gel surface next to one end of the IPG strip with forceps. Overlay the IPG strip and prestained markers with 3 mL of molten agarose gel overlay solution (see Note 57) and allow to set for 1 min. Insert the gels into the cassette carrier and add blank casset inserts to empty gel slots. Secure the gel cassets with the upper buffer chamber seal and place the assembly into the lower buffer chamber. Add approximately 4.0 L of 1× running buffer to the lower chamber and approximately 0.8 L of 2× running buffer to the upper chamber. Adjust the buffer levels of both chambers until they are equal and between the minimum and maximum fill lines marked on the tank (see Note 58). Attach the lid of the Electrophoresis Unit and connect to a suitable power supply. Run at 2.5 W per acrylamide gel for 30 min; 40 W for 1 h; and 80 W for 4–5 h (see Note 59).

3.6.5. Protein Detection and Gel Documentation 1. Carefully remove the IPG strips from the top of the gel and trim the gel bottom (see Note 60). 2. Submerge the gel in colloidal Coomassie™ stain and incubate overnight with gentle rocking (see Note 61). 3. Discard the stain and rock the gel in destain solution overnight. Repeat at least once with fresh destain solution until the background is clear. 4. Document gel with the GS-800™ calibrated imaging densitometer (Bio-Rad) or equivalent. The 2DE pattern of a representative S. pyogenes cell wall extract is presented in Fig. 2. The mutanolysin extracts are highly enriched with cell wall-associated proteins, as evidenced by the MALDI-TOF mass spectrometry identification of several well-characterized cell wall proteins of S. pyogenes (Table 1). 5. Store gel at 4 °C in a sealed plastic bag containing water (short term) or water with 0.005% (w/v) sodium azide for long term storage.

4. Notes 1. Dissolve the required amount of PMSF in a small volume of ice-cold isopropanol and adjust to final volume with chilled TES buffer. 2. APS breaks down in water resulting in rapid loss of reactivity. APS solutions should be prepared fresh each time. 3. Add required volume of 1 M DTT to buffer just before use.

2 3

1

Enolase (SEN) GAPDH Manganesedependent Superoxide dismutase

Protein

Virulence factor Virulence factor

Virulence factor

Function or pathway

Q5XDW3 Q8P0D4

P69950

Accession no.a

35.8 22.5

47.2

Molecular mass (kDa)b

b

Swiss-Prot/TrEMBL accession number. Theoretical values obtained from Swiss-Prot/TrEMBL database. c Number of tryptic peptides detected by MALDI-TOF MS that could be matched to the protein. d Percentage of protein sequence covered by the matched peptides.

a

Spot

5.34 4.87

4.74

pIb

19 9

24

57.0 57.0

68.9

Peptidematchc Coverage (%)d

Table 1 Landmark cell wall-associated proteins identiﬁed by MALDI-TOF peptide mass ﬁngerprinting analysis for S. pyogenes strain 5448 (Serotype M1)

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4. Bio-Lyte® 3/10 ampholyte (Bio-Rad) is recommended for all pH ranges. 5. Bromophenol Blue tracking dye allows monitoring of electrophoresis. 6. Combine ammonium sulfate, methanol and phosphoric acid. Bring to volume with Milli-Q® water and dissolve by vigorous stirring. Add the Coomassie™ G250 and stir vigorously with heating. Do not filter solution in order to retain colloidal dye particles. Discard stain after each use. 7. To minimize contamination and ensure cell viability, streak from glycerolized stocks of S. pyogenes stored at –80 °C rather than plates stored at 4 °C. Use fresh blood agar plates so that hemolysis is clearly visible. 8. Hold plate up to light or use a light box to ensure the colony is hemolytic. 9. The optical density at 600 nm (OD600 ) for late stationary phase cultures of S. pyogenes is approx 1.2. 10. PMSF irreversibly inhibits serine proteases and some cysteine proteases which may be liberated during the extraction process. 11. Seal the microcentrifuge tube lid with Parafilm® and secure to incubator in a horizontal position to ensure thorough mixing. 12. Mutanolysin cell wall extracts can be stored for at least 2 yr at –20 °C. 13. This step is used to verify the extraction of cell wall-associated proteins before sample preparation for 2DE. 14. The BCA assay is more sensitive than the Lowry assay and has less variability than the Bradford assay. Protein quantitation should be performed before 2DE as the solubilization solution reagents interfere with the BCA assay. Working Reagent is stable for one day at room temperature. 15. Standards should be prepared using the same diluent as the cell wall extracts to compensate for buffer interference. 16. To facilitate mixing, add each sample to the microtiter plate before adding the Working Reagent. 17. A purple colored BCA-copper complex is generated in the BCA protein assay. The samples may also be incubated at 60 °C for 15 min or at room temperature (25 °C) from 2 h to overnight. Incubation at higher temperatures will increase the sensitivity of the assay. 18. An absorbance range of 540–590 nm may also be used with minimal loss of signal. A standard curve should be determined for each assay. 19. Atmospheric oxygen inhibits the polymerization of acrylamide. For consistent results, degas the solution without the APS and TEMED under vacuum for at least 15 min. Add APS and TEMED and swirl the solution gently but thoroughly to minimize the introduction of oxygen. 20. The 12% resolving gel has an approximate separation range of 14.4 kDa to 120 kDa. 21. Remove the ethanol overlay after 1 h to prevent dehydration of the gel surface. 22. Slowly load the sample to avoid a diffuse loading zone and loss of band sharpness. Load 15 μL of 1× loading buffer in unused wells to prevent the lateral spread of adjoining samples.

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23. The voltage and current used for SDS-PAGE is potentially lethal. Use all equipment in accordance with the manufacturer’s instructions. 24. The detection limit for Coomassie™ Blue staining is 0.1–0.5 μg protein. 25. The Coomassie™ Blue stain can be reused several times. 26. The addition of a piece of sponge or paper towel to a corner of the tray will facilitate the removal of excess Coomassie™ Blue stain. Rapid destain solution can be recycled by filtering through activated carbon (Sigma). If loss of band intensity occurs because of excessive destaining, restain with Coomassie™ and destain as described. 27. The addition of glycerol to the final destain solution prevents the cracking of gels during drying. 28. TCA is a hazardous substance and care should be taken to avoid exposure. The TCA precipitation step is used to increase protein concentration before 2DE. 29. The pellet is usually invisible, but can sometimes be seen as a white smear. To avoid disturbing the pellet, carefully add the ethanol and gently tap the tube to mix. 30. Avoid over-drying as this will make the pellet difficult to resolubilize. TCA precipitated cell wall proteins can be stored for at least 2 yr at –20 °C. 31. Poor protein solubilization may cause horizontal or vertical streaking in 2D gels. Do not heat to resolubilize. Samples containing urea must not be heated above 35 °C to avoid protein carbamylation. 32. A water-bath sonicator is suitable for this step. Alternatively, a sonicator with a microtip may be used. 33. Centrifugation removes particulate material which can block the gel pores of the IPG strip, resulting in poor focusing and horizontal streaking. 34. The protocol may be stopped at this point and the sample stored at –20 °C. 35. Load up to 1 mg protein for Coomassie™ stained 2D gels. Sample overload may cause horizontal or vertical streaking. Do not exceed a total loading volume of 250 μL per strip. 36. A trace amount of Bromophenol Blue is included for monitoring of IPG strip rehydration and subsequent electrophoresis. 37. Evaporation will cause the urea to precipitate. 38. The minimum rehydration time is 11 h. 39. Thoroughly cleaned rehydration trays may be re-used. 40. Drainage of the oil removes unabsorbed protein and minimizes horizontal streaking in 2D gels. The IPG strip may be wrapped in plastic wrap and stored at –80 °C. 41. Always wear laboratory gloves when handling IPG strips and all apparatus/ solutions used in their preparation to prevent contamination from skin keratin. 42. Electrode wicks are highly recommended because they remove salts and other contaminants in the sample. 43. The majority of S. pyogenes cell wall-associated proteins have a pI between 4 and 7 (8). The appropriate pH range for other Gram-positive prokaryotes should be determined empirically.

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44. Covering the strip with mineral oil prevents evaporation and carbon dioxide absorption during focusing. 45. The maximum permissible voltage is 8,000 V. The focusing range for 11 cm IPG strips is 15,000–60,000 Vh. Incomplete or excessive isoelectric focusing may cause horizontal or vertical streaking in 2D gels. Focusing time should be kept to the minimum necessary and must be determined empirically. 46. If required, the IPG strip may be wrapped in plastic wrap and stored indefinitely at –80 °C without having a detrimental effect on the final 2D pattern. Thaw at room temperature for 10–15 min before equilibration. 47. Firmly push the separator sheets and cassettes against the bottom of the caster to ensure a good seal. If a full set of gels is not required, use blank cassette inserts with separator sheets to occupy the extra space. 48. Push down firmly during this step to ensure the faceplate forms a tight seal. 49. A homogenous gel containing 12.5% total acrylamide has a separation size range of 14 to 100 kDa. 50. For consistent results, degas the monomer solution (excluding the APS and TEMED) under vacuum for 15 min before adding the APS and TEMED. Degassing is not essential, but will accelerate polymerization. 51. For longer term storage, unload the gel cassettes from the caster, wrap in paper towel and completely submerge in gel storage buffer. Store in a sealed container at 4 °C for up to 3 months. 52. Wear gloves when preparing buffers and handling 2DE gels to prevent protein contamination, primarily with skin keratin. 53. IPG strip equilibration should be undertaken before second dimension separation. This step enhances the solubility of focused proteins and allows the binding of SDS for second dimension separation. Ineffective equilibration may cause vertical streaking in 2D gels. Thoroughly cleaned equilibration trays may be re-used. 54. Ensure the glass casset and gel surface are dry before loading the IPG strips. Remove excess water from the wells with blotting paper, taking care to avoid the gel surface. 55. This step lubricates the strip, preventing it from adhering to the glass plates during loading. 56. Two 11 cm strips or three 7 cm strips fit onto a 26 × 20 cm gel. Avoid air bubbles between the strip and the gel surface or between the strip backing and glass plate. A gap between strip and gel, or damage to the strip during application, may cause vertical streaking in 2D gels. 57. Melt the agarose gel overlay solution in a microwave oven. Allow to cool slightly (60 °C) to prevent the decomposition of urea in the equilibration buffer. Pipet slowly to avoid introducing air bubbles under or behind the strip, which may interfere with protein resolution. Air bubbles should be removed immediately with a spatula. 58. Depletion of ions in the running buffer can result in poor resolution and vertical streaking. To avoid this, discard the running buffer after approx 3 runs.

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59. Gels can be run overnight at 1.5 W per gel. The Bromophenol Blue in the agarose gel overlay solution is used to monitor migration. 60. Trim around the sides of the gel to ensure it does not adhere to the spacers and tear. 61. Colloidal Coomassie™ can detect down to 100 ng/protein spot.

Acknowledgments This work was supported by the National Health and Medical Research Council of Australia. We wish to thank Stuart Cordwell (School of Molecular and Microbial Biosciences, The University of Sydney, NSW, Australia) for assistance with the MALDI-TOF MS analysis. References 1. Ton-That, H., Marraffini, L. A., and Schneewind, O. (2004) Protein sorting to the cell wall envelope of Gram-positive bacteria. Biochim. Biophys. Acta 1694, 269–78. 2. van Heijenoort, J. (2001) Formation of the glycan chains in the synthesis of bacterial peptidoglycan. Glycobiology 11, 25R–36R. 3. Sjoquist, J., Movitz, J., Johansson, I. B., and Hjelm, H. (1972) Localization of protein A in the bacteria. Eur. J. Biochem. 30, 190–4. 4. Navarre, W. W., and Schneewind, O. (1999) Surface proteins of Gram-positive bacteria and mechanisms of their targeting to the cell wall envelope. Microbiol. Mol. Biol. Rev. 63, 174–229. 5. Chassy, B. M., and Giuffrida, A. (1980) Method for the lysis of Gram-positive, asporogenous bacteria with lysozyme. Appl. Environ. Microbiol. 39, 153–8. 6. Yokogawa, K., Kawata, S., Takemura, T., and Yoshimura, Y. (1975) Purification and properties of lytic enzymes from Streptomyces globisporus 1829. Agric. Biol. Chem. 39, 1533–43. 7. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–5. 8. Cole, J. N., Ramirez, R. D., Currie, B. J., Cordwell, S. J., Djordjevic, S. P., and Walker, M. J. (2005) Surface analyses and immune reactivities of major cell wall-associated proteins of group A Streptococcus. Infect. Immun. 73, 3137–46.

25 Cell Fractionation of Parasitic Protozoa Wanderley de Souza, José Andrés Morgado-Diaz, and Narcisa L. Cunha-e-Silva

Summary Cell fractionation, a methodological strategy for obtaining purified organelle preparations, has been applied successfully to parasitic protozoa by a number of researchers. These studies have provided new information of the cell biology of these parasites and have supported investigators to assume that some of the protozoa form the roots of the evolutionary tree of eukaryotic cells. The cell fractionation usually starts with disruption of the plasma membrane, using conditions that minimize damage to the membranes bounding intracellular organelles. An important requirement for successful cell fractionation is the evaluation of the isolation procedure that can be made by morphological and biochemical methods. The morphological approaches use light and electron microscopy of thin section of different fractions obtained, and the biochemical methods are based on the quantification of marker enzymes or other molecules (for instance, a special type of lipid, an antigen, etc.). Here we will present our experience in the isolation and characterization of some structures found in trypanosomatids and trichomonads.

Key Words: Cell fractionation; electron microscopy; marker enzymes; parasitic protozoa; trichomonads; trypanosomatids.

1. Introduction Parasitic protozoa are responsible for a large number of diseases affecting millions of people throughout the world. Some are caused by protozoa, which establish an initial binding to cells from the host and then invade them using several alternative mechanisms. Among these protozoa we can mention members of the Trypanosomatidae family, causative agents of Chagas’ disease and leishmaniasis, and members of the Apicomplexa group, which include From: Methods in Molecular Biology, vol. 425: 2D PAGE: Sample Preparation and Fractioation, Volume 2 Edited by: A. Posch © Humana Press, Totowa, NJ

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agents of malaria and toxoplasmosis. Other protozoa exert their parasitic activity via interaction with epithelial cells from the gastrointestinal and urogenital cavities. Examples include the agents of giardiasis, trichomoniasis, and amebiasis. To exert a parasitic activity these protozoa present several biosynthetic and metabolic pathways, which make them different from mammalian cells. Transmission electron microscopy of thin sections has provided most of the available information on the structural organization of parasitic protozoa. Structures typically found in mammalian cells, such as the nucleus, endoplasmic reticulum, Golgi complex, mitochondria and lysosomes are found in most, but not in all protozoa. Some lack structures such as the Golgi complex and mitochondria. Others present special secretory organelles, such as the micronemes, rhoptries, and dense granules found in Apicomplexa, or metabolic organelles, such as the hydrogenosome, found in trichomonads. One approach that has been used to obtain new information on structure and composition of structures and organelles found in parasitic protozoa is cell fractionation. Here we will present our experience in the isolation and characterization of some structures found in trypanosomatids and trichomonads. Figs. 1 and 2 show general views of the structural organization of Trypanosoma cruzi and Tritrichomonas foetus, respectively, based on information obtained by transmission electron microscopy. As can be seen in the figures these protozoa present well

Fig. 1. Longitudinal section of a Trypanosoma cruzi epimastigote depicting the the typical positions of the nucleus (N), kinetoplast (K), anterior flagellum (F) and reservosomes (R) at the posterior region of the parasite. Top inset shows a detail of reservosome structure and bottom inset shows Golgi complex (GC), always positioned next to the flagellar pocket, but that was not at the plane of the longitudinal section. Bars: 0.5 μm.

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Fig. 2. Longitudinal section of Tritrichomonas foetus showing the nucleus (N), Golgi complex (G), hydrogenosomes (H) and glycogen particles (asterisk) (Micrograph by M. Benchimol). Bar = 0.5 μm.

elaborated cytoskeletal structures, known as the subpellicular microtubules and the pelta-axostyle system, which make them resistant to cell breakage. 2. Materials 2.1. Cell Culture and Standard Equipment 1. Trypanosomatids: Liver Infusion Trypticase Medium (LIT) (1) supplemented with 10% heat-inactivated bovine serum (Gibco/BRL). 2. Trichomonads: Trypticase Yeast Maltose Medium (TYM) (2) supplemented with 10% heat-inactivated bovine serum (Gibco/BRL). 3. Ultrasonic apparatus (Sigma, GEX 600 Model) with standard probe (13 mm radiating diameter. 4. Sorvall RC 5B centrifuge with GSA and SS-34 Rotors.

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5. Ultracentrifuge with swinging-bucket (Beckman SW 50.1, SW 28 and fixed angle Type 65) rotors. 6. Potter-type homogenizer with Teflon pestle.

2.2. Electron Microscopy 1. Glutaraldehyde grade II (Ted Pella Inc.) is stored at 25% at –20 °C. Working solutions of 2.5% is prepared by diluting in Cacodylate buffer 0.1M, pH 7.2 (Sigma). 2. Osmium tetroxide (Ted Pella Inc.) is dissolved at 2% in distillated water and stored in a single use aliquot at –20 °C. Working solutions is prepared by dilution in cacodylate buffer plus 5 mM CaCl2 and 0.8% potassium ferrocyanide. 3. Acetone p.a: solutions of 30, 50, 70, 90, and 100%. 4. Epoxy resin: Polybed 812 (Electron Microscopy Sciences, EMS). 5. Aqueous solutions of uranyl acetate (Electron Microscopy Sciences, EMS) and lead citrate.

2.3. Isolation and Subfractionation of the Flagellum of Trypanosomatids 1. Buffer A: 25 mM Tris–HCl, pH 7.4, 0.2 mM EDTA, 5 mM MgCl2 , 12 mM μ-mercaptoethanol, 320 mM sucrose. 2. Sonication buffer: buffer A supplemented with 1% bovine serum albumin (BSA) and 1 mM CaCl2 and protease inhibitors cocktail (Sigma). 3. Sucrose gradient solutions for the isolation of flagellum: 0.8, 1.65, and 1.85M prepared in buffer A without sucrose. 4. 2% Triton X-100 prepared in buffer A. 5. Sucrose gradient to isolation of the flagellar membrane: 1.3, 1.6, 1.9, and 2.2M prepared in buffer A without sucrose. 6. 0.0001% trypsin (type XIII-treated, Sigma). 7. Soybean trypsin inhibitor (Sigma). 8. Continuous sucrose gradient ranging from 1.8 to 2.2M

2.4. Isolation of the Reservosome 1. TMS buffer: 20 mM Tris-HCl, pH 7.2, 2mM MgCl2 , 250 mM sucrose. 2. Sucrose solutions: 2.3, 1.2, 1.0, and 0.8 M prepared in TMS buffer without sucrose (TM buffer). 3. TMS buffer supplemented with a cocktail of protease inhibitors (Sigma).

2.5. Isolation of the Golgi Complex 1. Buffer G: 10 mM Tris-HCl, pH 7.2, 0.25M sucrose, 2 mM MgCl2 . 2. Hypotonic solution: G buffer without sucrose containing a cocktail of protease inhibitors.

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3. Gradient sucrose solutions: 2.3, 1.2, 1.0, and 0.8M prepared in G buffer without sucrose. 4. 150 mM sodium carbonate, pH 11.5.

2.6. Isolation and Subfractionation of the Hydrogenosomes 1. H buffer: 10 mM Tris-HCl, pH 7.2, 0.25M sucrose and 2 mM MgCl2 . 2. Percoll gradient solutions (Pharmacia): 53, 45, 24, and 18% prepared in G buffer. 3. V buffer: 2% Triton X-100, 50 mM 3-(N-morpholino)propanesulfonic acid (MOPS))-NaOH, pH 7.0, 0.1 mM MgCl2 , 1 nM dithiothreitol (Sigma) and 10% sucrose. 4. Proteinase K (Sigma) at a final concentration 0.5 mg/mL. 5. Quench solution: phenylmethylsulphonyl fluoride (PMSF) (final concentration 40 μg/mL).

3. Methods 3.1. Isolation of Trypanosomatid Structures 3.1.1. Isolation and Subfractionation of the Flagellum of Trypanosomatids Trypanosomatid flagellum exhibits a huge lattice-like filamentous structure called paraflagellar rod (PFR), running alongside a canonical 9+2 axoneme (3,4). The flagellar membrane, which has been considered a special domain, although continuous with cell body membrane, surrounds both structures. PFR plays a role in protozoa movement, as was indicated by a Trypanosoma brucei mutant that did not assemble the structure and was immotile (5). Flagellar membrane has a crucial participation in the interaction of T. cruzi with invertebrate host and parasite differentiation (6). The knowledge concerning the role played by trypanosomatid flagellum in the life cycle of these parasites and in the pathogenesis they cause in their vertebrate hosts has advanced using flagellar fractions as immunogen (7) or for biochemical and ultrastructural studies (8). There are two successful protocols for obtaining purified flagellar fractions: a classical one (9), deflagellating the protozoa and isolating the detached flagella by sucrose gradients, and a more simple method (10) that extracts the membranes of the whole protozoa and depolymerizes the cell body cytoskeleton by high salt treatment. The advantage of the latter method is that it does not take long, but it has a low yield, being limited to 105 parasites at a time. Here we describe the classical method that, although laborious, can be applied to 1012 parasites in a single experiment, yielding an abundant highly purified flagellar fraction. PFR is a highly conserved structure among trypanosomatids (11), allowing the adoption of the PFR from Herpetomonas megaseliae, a nonpathogenic

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trypanosomatid, as a model for ultrastructural and biochemical studies. The protocols presented below have already been published for whole flagella purification (9), PFR (12), and flagellar membrane isolation from both H. megaseliae and T. cruzi (8). 1. Start with 1011 –1012 protozoa. Harvest protozoa by centrifugation and wash twice in ice cold buffer A. All subsequent steps should be performed in an ice bath (see Note 1). 2. Washed protozoa (the pellet of the last washing step) are resuspended in sonication buffer: buffer A supplemented with 1% bovine serum albumin (BSA) and 1 mM CaCl2 and protease inhibitors cocktail. Cell density is very important in this step. If you started with 8 × 1011 protozoa, resuspend in 200 mL and sonicate in 50-mL aliquots (see Note 2). 3. We use an ultrasonic apparatus adjusted to 8% of maximum amplitude output. Typically, 6–8 sonication cycles of 10 s on and 5 s off are sufficient to deflagellate most cells (see Note 3). 4. Whole cells, deflagellated cell bodies and liberated nuclei are pelleted with a low speed centrifugation at 120g, 10 min (see Note 4). 5. Supernatants are centrifuged at 6,800g, 15 min, the pellet resuspended to 15 mL with buffer A and divided in six aliquots. 6. Each aliquot is equilibrated on top of 0.8M sucrose cushions and spun at 1,080g for 20 min (see Note 5). 7. Upper layer of each cushion is carefully removed and pelleted all together at 17,300g. This is the crude flagellar fraction (see Note 6). 8. The purification is achieved by laying 1-mL aliquots of crude flagellar fraction on top of a 1.65M/1.85M sucrose gradient, each step with 2 mL, and centrifuging at 130,000g for 3 h. 9. Flagella equilibrate at the interface 1.65M/1.85M that must be carefully removed with a long and thin Pasteur pipet. All the interfaces together are diluted with buffer A without sucrose and pelleted at 17,300g for 20 min. The pellet is resuspended in a small volume of buffer A (0.5 mL) and contains purified flagella. 10. Phase contrast microscopy can be used for initial examination of the flagellar fraction, but the only way of evaluating purity is electron microscopy. A small aliquot can be fixed by adding glutaraldehyde to the final concentration of 2.5% in buffer A and processed for electron microscopy (Fig. 3). (see Note 7). 11. H. megaseliae or T. cruzi flagella purified as above are submitted to detergent extraction by doubling sample volume with ice cold 2% Triton X-100 in buffer A and incubated for 15 min. on ice with intermittent agitation. 12. Sample is laid on top of a 1.3/1.6/1.9/2.2M sucrose and centrifuged at 130,000g for 3 h. 13. Purified flagellar membrane that equilibrates at the interface 1.3/1.6M is carefully removed, diluted with buffer A without sucrose, pelleted at 17,300g for 20 min, and resuspended in a small volume (0.05 mL) (Fig. 4).

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Fig. 3. Flagellar fractions from Herpetomonas megaseliae. Shows purified whole flagella, with preserved axonemes and PFRs enclosed by loose-fitting membrane profiles. Bar = 0.2 μm; Adapted from (8). 14. H. megaseliae or T. cruzi purified flagella are submitted to extensive membrane solubilization with three rounds of incubation in 2% Triton X-100 or Nonidet P-40 for 15 min at 4°C. After each round flagella are pelleted at 17,300g, 20 min. 15. Demembranated flagella are resuspended in buffer A and incubated very briefly (30 s) at 28°C with 0.001% trypsin (type XIII, TPCK-treated, Sigma). The reaction is stopped by addition of a 20-fold excess of specific trypsin inhibitor (soybean trypsin inhibitor, Sigma). Shake in a Vortex at maximum speed for 1

Fig. 4. Flagellar fractions from Herpetomonas megaseliae. Shows flagellar membranes purified from Fig. 3 preparation. Bar = 0.2 μm; Adapted from (8).

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min, divide in six aliquots and apply on a continuous sucrose gradient ranging from 1.8 to 2.2M. 16. Centrifuge at 130,000g for 3 h. Collect 0.8-mL aliquots from top to bottom, dilute with buffer A without sucrose and pellet at 17,300g for 30 min. The third and fourth fractions from top to bottom are highly purified PFRs, as can be evaluated by electron microscopy (Fig. 5) and by using antibodies against major PFR proteins (Fig. 6B) and alpha-tubulin (Fig. 6C) in Western blots. SDS-PAGE analysis of purified PFRs (Fig. 6A) allowed for the first time the identification of some minor proteins that constitute this complex structure.

3.1.2. Isolation of the Reservosome Reservosomes are storage organelles from T. cruzi epimastigotes that concentrate proteases and are essential for differentiation into trypomastigote forms. They were considered prelysosomes but the lack of a molecular marker has precluded a more detailed studied of their role in T. cruzi life cycle. Their purification (13) opened up the possibility of defining the necessary marker. 1. Start with 3 × 1010 epimastigotes from mid log phase cultures (see Note 8). 2. Harvest protozoa by centrifugation at 4,800g, 10 min, 4°C and wash in cold TMS buffer (see Note 9). 3. Resuspend in 45 mL of TMS (about 6.5 × 108 parasites/mL) and divide in three aliquots. 4. Sonicate each aliquot at 10% of maximum amplitude of ultrasonic apparatus, in about 15 cycles of 2 s on and 1 s off. (see Note 3). 5. Centrifuge at 2,450g, 10 min, pour the supernatant carefully into another tube and discard the pellet, containing non-ruptured cells, nuclei, kinetoplasts etc.

Fig. 5. Flagellar fractions from Herpetomonas megaseliae. Shows that PFRs remain organized after purification from demembranated H. megaseliae purified flagella. Bar = 0.1 μm; Adapted from (12).

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Fig. 6. (A) SDS-PAGE gel of fractions from the PFR purification gradient. Lanes 1–4 represent fractions 3–6, in this order. Besides the major PFR proteins (arrowhead), fractions 3 and 4 are clearly enriched in a number of minor bands whose positions and estimated molecular masses are indicated on the left. Molecular mass markers are indicated on the right. (B and C) Western blots of different steps and gradient fractions from the paraflagellar rod (PFR) purification procedure probed with anti PFR major proteins antibody (panel B), and antialpha-tubulin monoclonal antibody (panel C). In both panels lanes 1–4 represent fractions 3–6, in this order, from the PFR purification gradient. Molecular mass markers are indicated on the left. Adapted from (12).

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6. Mix the supernatant with an equal volume of 2.3M sucrose in TMS and divide in two Beckman SW28 centrifuge tubes (usually 12 mL in each tube) (see Note 10). 7. Repeating the procedure with the other two aliquots, all six tubes of the rotor will be used. 8. Overlay carefully with 10 mL of 1.2M sucrose in TMS, 9 mL of 1.0M sucrose and 8 mL of 0.8M sucrose. 9. Centrifuge at 97,000g for 150 min, without brake. 10. Collect the interfaces 0.8M/1.0M (B1) and 1.0M/1.2M (B2), 1.2M/1.27M (B3) and the pellet (P), dilute with TM (TMS without sucrose) and pellet at 90,000g for 20 min. Reservosomes are highly purified in B1 (Fig. 7) and very enriched in B2. Microsomes and glycosomes are the major components of B3. Acidocalcisomes are enriched in the pellet. Fig 7 here 11. A small aliquot of each fraction is fixed by adding glutaraldehyde directly in TMS for electron microscopy (see Note 7), another aliquot is reserved for marker enzyme assays (see next step), and protease inhibitors are added to a third aliquot for SDS-PAGE and Western blot assays. Purified fractions must be stored below –70°C. 12. To evaluate the purity of the fractions marker enzymes for the organelles that could be contaminants of reservosome fraction should be assayed: succinate cytochrome c reductase for mitochondria (14), pyrophosphatase for acidocalcisomes (Table 1) and hexokinase for glycosomes (15).

Fig. 7. Trypanosoma cruzi reservosomes purified fraction. Adapted from (13). Bar = 1 μm.

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3.1.3. Isolation of the Golgi Complex In parasitic protozoan the Golgi complex is much less well organized, less prominent, and correspondingly more difficult to isolate than in mammalian cell. The success of the organelle isolation is dependent on choosing a good cell disruption method, which is dependent on the morphological characteristics of the parasite. Morphological studies have shown that the Golgi of trypanosomatids, and in particular of Trypanosoma cruzi, is made up of 4–10 stacked cisternae localized at the anterior region of the cell, close to the flagellar pocket (16). A method that preserves the Golgi with minimum damage during cell disruption was developed resulting in a fraction highly enriched in stacked Golgi cisternae and vesicles (19), as determined by electron microscopy (Fig. 8), and also in galactosyltransferase, O-- GlcNA transferase and acid phosphatase activities. Contamination with other organelles (endoplasmic reticulum, plasma membrane, mitochondria, and glycosomes) is minimal, as analyzed by their respective marker enzymes (Table 2). All centrifugation steps and other operations are performed at 4 °C or in an ice bath. 1. Harvest cells in a GSA Rotor at 1,000g for 10 min and wash three times in G buffer. 2. Resuspend the cells in a hypotonic solution (G buffer without sucrose) containing a cocktail of protease inhibitors and incubated in this solution for 10 min. 3. Homogenize the cells by sonication on ice with 15 cycles of 2 s with 1 s rest between cycles in an ultrasonic apparatus. Monitor the cell disruption by phasecontrast microscopy (see Note 3). 4. Immediately add to the homogenate a concentrated sucrose solution, prepared in G buffer to a final concentration of 0.25M sucrose to minimize osmotic damage.

Table 1 Enzyme assays in the epimastigote subcellular fractions of the puriﬁcation of reservosome of Trypanosoma cruzi

Total homogenate Supernatant from 2,450g B1 B2 B3 B4 a b

Succinate cytochrome c reductasea

Pyrophosphataseb

0.030 0.043 0 0.006 0.044 0.830

0.11 0.14 0 0.02 0 0.15

Specific activity in mM min−1 mg protein−1 . Specific activity in μM pyrophosphate consumed min−1 mg protein−1 .

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Fig. 8. General view of the Golgi fraction of Trypanosoma cruzi showing Golgi complex cisternae and several Golgi cisternae profiles. After (19). Bar = 0.5 μm.

5. Centrifuge at 2,500g for 15 min and discard the pellet containing unbroken cells, nuclei, and kinetoplasts.

Table 2 Enzyme activities in Golgi fraction and other subcellular fractions of Trypanosoma cruzi.a Enzyme Activitybc Assay NADPH cyt c reductaseb Acid phosphataseb 5’-nucleotidaseb Succinate cyt c reductaseb Hexokinaseb O-a-GlcNAc transferasec Gal transferasec

Homogenate fraction 025 30 0101 007 0032 0076 128

PNS 034 45 0157 004 00436 0248 656

GF 0043 152 006 003 00034 2044 11480

PNS, post-nuclear supernatant; GF, Golgi fraction The results are of a typical experiment in which each number represents the means of duplicate determinations. Minor modifications were observed in at least three independent experiments. b Specific activity expressed as μM of product released min−1 mg protein−1 . c Specific activity expressed as pM of [3 H]GlcNAc or [3 H]Gal transferred h−1 mg protein−1 . a

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6. Harvest the postnuclear supernatant (PNS), add an equal volume of 2.3M sucrose and homogenize for 10 min. 7. Load 15 mL of PNS onto the bottom of Beckman SW 28 Rotor ultracentrifuge tubes and overlay sequentially with 7 mL each of 1.2M, 1.0M, and 0.8M sucrose. 8. Centrifuge the gradient at 95,000 x g for 1h 30 min. The band at the interface of 1.0/1.2M sucrose corresponds to highly enriched Golgi complex fraction. 9. Collect the fraction by diluting with G buffer without sucrose and centrifuging at 80,000g for 45 min (Beckman Type 65 Rotor).

3.2. Isolation of Trichomonad Structures 3.2.1. Isolation of the Golgi Complex Tritrichomonas foetus, a member of the Trichomonadidae family, possesses a well developed Golgi complex, consisting of tightly packed elongated cisternae often with dilated terminal regions, localized at the anterior portion of the cell and in close association with the parabasal filament (18). The following protocol yields two subfractions containing light (GF1) and heavy (GF2) Golgi membranes (17). At the ultrastructural level GF1 fraction is constituted only of smooth-membrane structures, particularly cisternae and secretory vesicles (Fig. 9A) and contains a 20-fold enrichment of galactosyltransferase activity. GF2 fraction is composed of profiles of stacked Golgi membranes with three or more tightly apposed cisternae with dilated borders and associated to vesicles (Fig. 9B). This latter fraction contains a 7-fold enrichment in galactosyltransferase activity. Minimal contamination with other organelles, such as endoplasmic reticulum, hydrogenosomes, and plasma membrane is observed in these fractions, as quantified by their respective enzymatic markers (Table 3). All centrifugation steps and other operations are performed at 4 °C or in an ice bath. 1. Harvest the cells in a GSA Rotor at 1,000g for 10 min and wash three times in G buffer. 2. Resuspend the cells in G buffer containing a cocktail of protease inhibitors and homogenize using 180 strokes of a Potter-type homogenizer. Stop the homogenization procedure when about 90% cell breakage is achieved as ascertained by phase-contrast microscopy. 3. Centrifuge at 2,500g for 10 min to discard the pellet containing unbroken cells, nuclei, and hydrogenosomes. 4. Add to the supernatant (PNS) an equal volume of 2.3M sucrose to make a 1.4M sucrose final concentration and homogenize for 10 min. 5. Load 15 mL of PNS onto the bottom of SW28 ultracentrifuge tubes and overlay in succession with 7 mL each of 1.2M, 1.0M, and 0.8M sucrose. 6. Centrifuge at 95,000g for 2.5 h and carefully remove two bands from the top at 0.8/1.0 (GF1) and 1.0/1.2M (GF2).

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Fig. 9. General aspect of the Golgi fractions of Tritrichomonas foetus. (A) GF1 fraction showing several vesicles (arrowheads) and cisternae profiles (arrows). (B) GF2 fraction showing several intact stacks of Golgi complex (arrowheads). After (17). Bars = 0.2 μm (A) and 0.5 μm (B).

7. Dilute these fractions in G buffer without sucrose and collect them by centrifugation at 80,000g for 30 min (Beckman Type 65 Rotor). 8. Further fractionation of these two compartments can be attempted by alkaline treatment to obtain the Golgi content and Golgi membrane subfraction. Blend the

Table 3 Distribution of Galactosyltransferase and other enzyme markers in Golgi fractions of Tritrichomonas foetus.a Fractions

Homogenate PNS GF2 GF1

Total Galactosyl NADPH cit c NADP-malic 5’Nucleotidase protein transferase reductase enzyme (mg) (pmole/30 (μmole/min/mg (μmole/min/mg min/mg protein) protein) protein) 3920 1040 20 10

4434 6804 31494 87400

0035 00079 00029 n.d.

0012 0026 0006 n.d.

0.93 0.89 1.50 0.66

a Results are expressed as the means of duplicate determinations. n.d., not detectable; Number of experiments is given in parenthesis; PNS, postnuclear supernatant; GF2, heavy Golgi fraction; GF1, light Golgi fraction.

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Golgi fractions (GF1 and GF2), collect by centrifugation for 30 min at 80,000g (Beckman Type 65 Rotor), and treat with 150 mM sodium carbonate, pH 11.5, for 30 min. 9. Collect the subfractions, again as in step 7.

3.2.2. Isolation and Subfractionation of the Hydrogenosome T. foetus is a facultative anaerobic protist that lacks mitochondria, but contains another organelle, designated as the hydrogenosome. This organelle appears as a spherical or slightly elongated structure with a mean diameter of 0.5 μm, reaching 2 μm in dividing organelles. A common characteristic of most hydrogenosomes is the presence of a peripheral vesicle, which has been suggested to be a specialized sub-compartment (20). Additional information on this important organelle, which seems to be the main target of drugs used in the chemotherapy of trichomoniasis, can be obtained using hydrogenosomes isolated in a way that preserves their structure. Using Percoll gradients, we developed a procedure (21), which yielded the purest fraction obtained for hydrogenosomes of T. foetus (Fig. 10A) and allowed further sub-fractionation to isolate the peripheral vesicle (Fig. 10B). It is enriched about 12-fold in malic enzyme activity and free of other contaminant organelles (Table 4). Major hydrogenosome proteins were identified by SDS-PAGE (Fig. 10C). All centrifugation steps and other operations are performed at 4 °C or in an ice bath. 1. Harvest the cells in a GSA Rotor at 1,000g for 10 min and wash three times in H buffer. 2. Resuspend the cells in 30 mL of H buffer and homogenize using about 180 strokes of a Potter-type homogenizer. Stop the homogenization while some unbroken cells are still present. 3. Dilute the homogenate in the proportion 1:10 with H buffer and centrifuge at 1,500g for 10 min. Discard the pellet containing unbroken cells, nuclei and costa. 4. Centrifuge the supernatant at 4,000g for 10 min. The pellet of this centrifugation corresponds to an enriched hydrogenosomal fraction. 5. Resuspend the pellet in ice-cold H buffer and layer aliquots of 1 mL on the top of a Percoll gradient of 53, 45, 24, and 18% in H buffer (1 mL each solution). 6. Centrifuge the gradient using a swinging bucket rotor (Beckman SW 50.1) at 36,000g for 30 min, without using the brake. The pure fraction containing wellpreserved hydrogenosomes is recovered from the pellet of the Percoll gradient. 7. Incubate a sample of the purified hydrogenosomal fraction in V buffer on ice for 1 h. and centrifuge the suspension at 25,000g for 20 min (Beckman Type 65 Rotor). The supernatant (TX-supernatant) is made up of solubilized hydrogenosome membranes and the pellet (TX-pellet) of hydrogenosomal matrix attached to the peripheral vesicle.

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Fig. 10. (A) Electron micrograph showing the purified hydrogenosomes of Tritrichomonas foetus. The hydrogenosome matrix is homogeneous and the peripheral vesicles present an electron-dense content. Bar = 1 μm (B) Purified fraction of hydrogenosomal flat vesicles Bar = 0.5 μm (C) SDS-PAGE of the fractions obtained during the fractionation of T. foetus. Lane 1: whole homogenate, Lane 2: fraction before the Percoll gradient, Lane 3: purified hydrogenosomes. After (21).

Table 4 Speciﬁc activity and recovery of malic dehydrogenase in fractions of the puriﬁcation of hydrogenosomes of Tritrichomonas foetus.a Fractions Whole homogenate Po fraction Pellet gradient

Total protein (mg)

Specific activity (μmol min−1 mg protein)

4500 580 80

0035 0147 0421

Enrichment Yield (%) (-fold) 10 42 120

100 54 22

a Po is the fraction before the Percoll gradient. Results are expressed as the means of duplicate determinations in three representative experiments.

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8. Wash a sample of the TX-pellet twice in H buffer and incubate for 45 min at room temperature with proteinase K (final concentration 0.5 mg/mL). 9. Quench the mixture with PMSF (final concentration 40 μg/mL) on ice, for 15 min and centrifuge at 350,000g for 30 min (Beckman Type 65 Rotor). The pellet of this centrifugation corresponds to the purified peripheral vesicle.

4. Notes 1. Three liters of axenic culture of H. megaseliae promastigotes in Warren’s medium supplemented with 5% fetal calf serum are enough for obtaining 8 × 1011 protozoa, after 30–36 h at 28°C. The same cell density of T. cruzi epimastigotes can be obtained with 5 L of parasites cultivated for 5 d at 28°C in LIT medium supplemented with 10% fetal calf medium (1). A good rotor to harvest a large volume culture is GSA of Sorvall ultracentrifuges. 2. It is not necessary to use purified BSA to prepare sonication buffer; fraction V grade is good enough. The composition of protease inhibitors cocktail may vary, it is very important that it contains an irreversible cysteine protease inhibitor, such as E-64, because this class of protease is the most active in trypanosomatids. We use protease inhibitors cocktail for general use, from Sigma. 3. It is essential to control deflagellation by phase contrast microscopy and stop the procedure when the majority, but not all the protozoa are deflagellated, otherwise some nuclei may rupture and free chromatin aggregates the organelles, including free flagella. 4. Turn off centrifuge brake because this loose pellet resuspends easily. This step and the next are performed with Sorvall SS-34 fixed angle rotor. The supernatant of the low speed centrifugation should be controlled by phase contrast microscopy and this step should be repeated twice to clean the supernatant from whole cells and deflagellated cell bodies. 5. The sucrose cushion must be spun in a swinging bucket rotor, as Beckman SW50.1, which must have each tube filled up to total volume (5 mL). In this step as well as in the gradient, the rotor should stop without using the brake. 6. To improve the isolation yield, the cushion can be repeated using the bottom layer as the starting sample of step 6. 7. A brief protocol for sample processing for electron microscopy: as fractions are easy to fix, 30 min of incubation in fixative solution is enough. On the other hand, fractions are not easy to wash, as the usual centrifugation of few minutes in a microfuge may not be sufficient to pellet the sample adequately and thus you would be losing your sample in the washing steps. To avoid this, wash in the ultracentrifuge at the same speed used to obtain the fraction until before osmium incubation when the fraction becomes denser. If you cannot process the samples in a short time (next day), change the fixative solution to 2.5% glutaraldehyde in 0.1 M cacodylate buffer pH 7.2, as Tris buffers are not good for electron microscopy. After fixation, samples are washed twice in 0.1M cacodylate buffer and post fixed with 1% osmium tetroxide in 0.1M cacodylate buffer containing

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0.8% potassium ferrocyanide and 5 mM calcium chloride for 20 min at room temperature in the dark (wear gloves and work in a fume hood!). Wash three times (or until the supernatant is clear) in 0.1M cacodylate buffer. Dehydrate in acetone series from 30 to 100%, 10 min incubation in each bath, repeating the last one twice. Infiltrate slowly in Epon (acetone + Epon 1:1 overnight, Epon for 6 h). Embed in Epon at 60o C for 2 d. 8. Epimastigotes from 5-day-old cultures are ideal for reservosome purification. Three liters of culture in LIT medium supplemented with 10% fetal calf serum (1) maintained at 28°C under gentle orbital agitation are enough to obtain 3 × 1010 parasites. 9. It is very important to work fast and in an ice bath, because reservosomes start to degrade their contents as soon as parasites are washed out from culture medium. It is adequate to add protease inhibitors to TMS buffer (mainly E-64, inhibitor of cysteine proteases, highly concentrated in the organelle), unless you just want to study the properties of reservosome proteases and their inhibitors. 10. The bottom step of this gradient contains the sample and corresponds to 1.27M sucrose, very close in density to the next step. So, take care when preparing and measuring volumes of sucrose solutions before mixing sample with 2.3M sucrose.

Acknowledgments The authors dedicate this chapter to the memory of Luiz Henrique Monteiro Leal, who greatly contributed to improve the methods of cell fractionation of parasitic protozoa, and passed away during the elaboration of this manuscript. References 1. Camargo, E. P. (1964) Growth and differentiation in Trypanosoma cruzi. I. Origin of metacyclic trypanosomes in liquid media. Rev. Inst. Med. Trop. Sao Paulo 6, 93–100. 2. Diamond, L. S. (1957) The establishment of various trichomonads of animals and man in axenix cultures. J. Parasitol. 43, 458–90. 3. Fuge, H. (1969) Electron microscopic studies on the intra-flagellar structures of trypanosomes. J. Protozool. 16, 460–6. 4. Farina, M., Attias, M., Souto-Padrón, T., and De Souza, W. (1986) Further studies on the organization of the paraxial rod of trypanosomatids. J. Protozool. 33, 552–7. 5. Bastin, P., Sherwin, T., and Gull, K. (1998) Paraflagellar rod is vital for trypanosome motility. Nature 391, 548–548. 6. De Souza, W. (1984) Cell biology of Trypanosoma cruzi. Int. Rev. Cytol. 86, 197–285. 7. Ruiz, A. M., Esteva, M., Riarte, A., Subias, E., and Segura, E.L. (1986) Immunoprotection of mice against Trypanosoma cruzi with a lyophilized flagellar fraction of the parasite plus adjuvant. Immunol Lett. 12, 1–4.

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8. Cunha-e-Silva, N. L., Hasson-Voloch, A., and De Souza, W. (1989) Isolation and characterization of a highly purified flagellar membrane fraction from trypanosomatids. Mol. Biochem. Parasitol. 37, 129–36. 9. Cunha, N. L., De Souza, W., and Hasson-Voloch, A. (1984) Isolation of the flagellum and characterization of the paraxial structure of Herpetomonas megaseliae. J. Submicrosc. Cytol. 16, 705–13. 10. Robinson, D., Beattie, P., Sherwin, T., and Gull, K. (1991) Microtubules, tubulin, and microtubule-associated proteins of trypanosomes. Methods Enzymol. 196, 285–99. 11. Gadelha, C., Lebowitz, J. H., Manning, J., Seebeck, T., and Gull, K. (2004) Relationships between the major kinetoplastid paraflagellar rod proteins: a consolidating nomenclature. Mol. Biochem. Parasitol. 136, 113–115. 12. Moreira-Leite, F.F., De Souza, W. and Cunha-e-Silva, N.L. (1999) Purification of the paraflagellar rod of the trypanosomatid Herpetomonas megaseliae and identification of some of its minor components. Mol. Biochem. Parasitol. 104, 131–40. 13. Cunha-e-Silva, N. L., Atella, G. C., Porto-Carreiro, I. A., Morgado-Diaz, J. A., Pereira, M. G., and De Souza, W. (2002) Isolation and characterization of a reservosome fraction from Trypanosoma cruzi. FEMS Microbiol. Lett. 214, 7– 12. 14. King, T. E. (1967) The Keilin-Hartree heart-muscle preparation. Methods Enzymol. 10, 202–8. 15. Rodrigues, C. O., Scott, D. A., and Docampo, R. (1999) Characterization of a vacuolar pyrophosphatase in Trypanosoma brucei and its localization to acidocalcisomes. Mol. Cell. Biol. 19, 7712–23. 16. Figueiredo, R. C. B. Q. and Soares, M. J. (1995) The Golgi complex of Trypanosoma cruzi epimastigote forms. J. Submic. Cytol. Pathol. 27, 209–15. 17. Morgado-Díaz, J. A., Monteiro-Leal, L. H., and De Souza, W. (1996) Tritrichomonas foetus: Isolation and Characterization of the Golgi Complex. Exp. Parasitol. 83, 174–183. 18. Benchimol, M., Ribeiro, K. C., Mariante, R. M., and Alderete, J. F. (2001) Structure and division of the Golgi complex in Trichomonas vaginalis and Tritrichomonas foetus. Eur. J. Cell Biol. 80, 593–607. 19. Morgado-Díaz, J. A., Nakamura, C. V., Agrellos, O. A. A., et al. (2001) Isolation and characterization of the Golgi complex of the protozoan Trypanosoma cruzi. Parasitology 123, 33–43. 20. Benchimol, M., Almeida, J. C. A., and De Souza, W. (1996) Further studies on the organization of the hydrogenosomes in Tritrichomonas foetus. Tiss. Cell. 28, 287– 299 21. Morgado-Díaz, J. A. and De Souza, W. (1997) Purification and biochemical characterization of the hydrogenosomes of the flagellate protozoan Tritrichomonas foetus. Eur. J. Cell Biol. 74, 85–91.

Index A Acrylamide-Bis solution, 205 Affinity Depletion Cartridge for removal of HSA, 91 Albumin and immunoglobulin-depleted plasma proteins 1D gel electrophoresis assessment of fractions, 20–21 UV absorbance assessment of fractions, 21–22 Albumin (BSA) standards color response curve using BCA assay, 123 Albumin-depleted plasma proteins 1D gel electrophoresis assessment of fractions, 18–19 UV absorbance assessment of fractions, 19–20 Alkaline phosphatase assay, biochemical principle, 252, 254 Anti-HSA POROS cartridge, 23 Anti-human albumin monoclonal antibody, 15 Arabidopsis thaliana chloroplasts, isolation and preparation, 171–172 chloroplast isolation, 174–176, 178–181 establishing yield and intactness of chloroplasts, 176 growth of Arabidopsis seedlings, 173–174, 177 preparation for proteomics, 176–177, 182–183 B BCA™ Protein Assay Kit, 117 Beconase AQ pump aspirator spray device, 79, 80

Bicinchoninic acid protein assay, 30, 121, 297 Biological Variation Analysis mode (BVA), 11 Bovine serum albumin (BSA), 204 Bradford protein assay, 121 Bronchoalveolar lavage fluid (BALF) cellular pattern of, 72–73 2-DE analysis, 71–72 during fiber-optic bronchoscopy, 67 proteome analysis for, 68 sample preparing dialysis-lyophilization, 71 ultra filtration, 71 C C. albicans protoplasts in active cell wall regeneration, proteomics of proteins secreted from, 261 Castor bean endosperm isolation and fractionation of ER for proteomic analysis, 203–214 isolation of, 206–208 mass spectrometer compatible silver staining, 205, 209–211 2D gel protein profiling of, 205–206, 211–213 SDS-PAGE analysis of developing and germinating of, 208–209 SDS-PAGE analysis of purified ER, 204–205 tissue homogenization and ER purification, 204 Cell physiology and disease molecular study, 139 cell culture and lysis, 142 isolation of nickel-binding proteins, 142, 143–144

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334 preparing samples for isolation of nickel-binding proteins, 143 silver staining of metal-affinity enriched proteins compatible for mass spectrometry (MS), 142–144 Cell wall incorporation of GPI-anchored proteins model, 232 Cell wall proteins (CWP), 187 analysis and extraction protocol by bioinformatics, 198 separation and identification of protein, 197–198 extraction and analysis from liquid culture medium of seedlings, 189 Cellular fungi sample preparation procedure, 261–266 cell culture, 267 cell disruption and protein solubilization, 267, 268–270 S-35 in vitro cell labeling, 267–268 scintillation counting and protein assay, 268, 270 Cerebrospinal fluid (CSF) LC/MS/MS analysis processing, 63 processing for 2-Dimensional gel electrophoresis, 59–60 proteomics, 53–56 gel electrophoresis, 57 LC-MS/MS mass spectrometry, 58 MALDI-TOF mass spectrometry, 58 processing for 2-dimensional gel electrophoresis, 59–60 processing for LC/MS/MS analysis, 57–58 silver staining and in-gel digestion, 57 simplification by removal of abundant proteins, 58–59

Index Chicken IgY antibodies, immunoaffinity fractionation of plasma proteins IgY-12 and LC2/ LC10 high capacity spin column kit, 43 IgY-12 microbeads for 96-well filter plates, 43 SDS gel electrophoresis, 43–44 Chloramphenicol, 224 Chloroplast isolation buffer (CIB), 174 isolation procedure, 175 light and electron micrographs, 180 lysis and solubilization, 182 protein samples by DIGE 2D-PAGE, 182 proteomic studies, 173 CLUSTAW program in PIR, 84 Collision-induced dissociation (CID), 136 Cysteine-specific protein labeling using CyDye DIGE fluor saturation dyes, 2 Cytoplasmatic proteins buffers for western blot analysis, 103 cell culture and wash buffer, 102 cell harvest, 104 comparison of 2DE pattern with pattern of whole cell protein, 109 coomassie brilliant blue staining solutions, 104 isolation from cultured cells 2-D gel electrophoresis, 101, 107–108 buffers and reagents for, 102–103 precipitation of cytoplasmatic proteins, 104–105 silver staining solutions, 103 western blot analysis for detection of purity, 105–107 D DeCyder software, 10–11 Depleted protein samples TCA precipitation for 2D gel electrophoresis, 22–23

Index Differential in-gel analysis (DIA), 11 DIGE saturation labeling conditions, 7 DNA electrophoresis, 156 DryStrip Reswilling tray, 70 Dulbecco’s Phosphate Buffered Saline (D-PBS), 116 E Econo-Pac® 10DG desalting column, 190 Electrospray quadrupole time of flight mass spectrometer, 80, 83 Eosin stain, 133 Epithelium lining fluid (ELF), 67 S-Ethanol-cysteine residue, 63 Etioplasts, dark-grown plants, 173 F FFPE tissue, 137 Filamentous fungi secreted proteins isolation and enrichment, 274–284 centrifugation and ultrafiltration, 277 enzymatic and chemical deglycosylation, 277–278 filtration and lyophilization, 277 isolation and concentration of supernatant broth, 278–280 precipitation with trichloroacetic acid, methanol and chloroform, 277 SDS-PAGE, 278, 282, 301 two-dimensional gel electrophoresis, 278, 282 Fluorescence dye labeling optimization, 6–7 Fluorescence resonance energy transfer (FRET), 4 Foetal bovine serum (FBS), 116 Formalin-fixed, paraffin-embedded (FFPE) tissues, 132 sample preparation for mass spectrometry analysis, 131 laser-capture microdissection, 134 LC-MS/MS analysis of tryptic peptides, 133–134

335 mass spectrometry analysis and bioinformatic analysis, 136 nanoflow RPLC-MS/MS analysis, 135–136 protein extraction and trypsin digest, 133–135 tissue processing, 133, 134 trypsin digest, 133 trypsin-mediated 18 O-labeling, 135 G Gamborg liquid medium, 190 Gel pieces, reduction and alkylation, 61–62 Glycosyl phosphatidylinositol (GPI)-cell wall proteins, 220 Gram-positive bacterial cell wall-associated proteins isolation and solubilization bacterial culture, 296, 299 BCA protein assay, 300 casting 2D gels, 304 cell wall-associated proteins extractions, 296–297, 300 first dimension IEF, 303–304 protein detection and gel documentation, 306 SDS-polyacrylamide gel electrophoresis (SDS-PAGE), 297 second dimension SDS-PAGE, 304–306 trichloroacetic acid (TCA) precipitation, 298, 302 Green plant tissue protein extraction, 149–152 H Hematoxylin stained histological classification, 5 Herpetomonas megaseliae, 319, 320 HiLoad 16/60 Superdex 75 prep grade column, 91

336 Hi-Trap SP equilibration buffer, 190 HONE1 cells, Western blot analysis of cytoplasmatic fraction, 107 Human BALF proteins, 2-D gel electrophoresis, 69 Human nickel allergy, 140 Human plasma albumin and immunoglobulin depletion of, 15–19 proteins by 1D gel electrophoresis, 20–21 albumin depletion, 16–17, 18 albumin-depleted plasma proteins, 18–20 blood sampling, 16 chromatography of immunoaffinity separation of using IgY-12 high capacity LC2 column, 46 1D and 2D gel Electrophoresis, 17 immunoglobulin depletion, 17, 20 samples collection of, 17–18 TCA precipitation of proteins, 17 Human serum chromatogram for affinity removal of high-abundant proteins from, 31 2D gel electrophoresis of, 34 1D SDS gel electrophoresis of, 33 high-abundant proteins, immunoaffinity depletion of, 29–30 immunodepleted serum and bund fraction processing, 30 low-abundant proteins, reversed-phase separation of, 30, 35–36 multi-component immunoaffinity subtraction and reversed-phase chromatography, 27–39 proteins composition, 28 I IgY-12 Microbeads regeneration, 45 ImageQuant™ software, 11 Iminodiacetic acid (IDA), 140

Index Immobiline Dry Strip™ (IPG) kit, 205 Immobiline Dry Strip Reswelling Tray™ , 205 Immobiline™ DryStrip Kit, 118 Immobilized metal ion affinity chromatography (IMAC), 139 Immobilized pH gradient, 70 Immunoaffinity subtraction (IAS), 94 Immunoglobulin depletion, 20 In-gel digestion procedure, 61–62 Inter simple sequence repeat PCR (ISSR-PCR) technique, 156 IPG strips, 71–72 IPG-strip rehydration buffer, 22–23 Isoelectric focusing (IEF) tube gels, 9 K Keratin proteome in normal NLF, 84 Kinematica Model PT10-35, 174 Knexus program, 63 L Labile CWP analysis, 191 Labile /weakly bound CWP extraction by nondestructive techniques cell suspension cultures, 194 liquid culture medium of seedlings, 193–194 Laemmli sample buffer, 17 LC-MS/MS mass spectrometry, 63–64 Liquid chromatography-coupled electrospray ionisation MS (LC-ESI-MS), 12 Liquid Tissue™ -MS protein prep kit, 133 Liver Infusion Trypticase Medium (LIT), 315 Lowry protein assay, 80 LTQ ion trap mass spectrometer, 64 Lumbar puncture (LP), 54 Lysis buffers, 176

Index M Madin-darby canine kidney (MDCK) cells sample preparation from culture medium, 113–128 cell culture, 114–116, 119 2-D gel electrophoresis of, 115 protein quantitation by BCA assay, 117, 121–122 rehydration loading and isoelectric focusing, 117–118, 122–125 ultracentrifugation and ultrafiltration, 116–117, 119–120 MALDI-TOF spectrometry, 58, 63 MASCOT MS/MS ion search software, 84 Mayer’s hematoxylin stain, 133 Metal oxide affinity chromatography (MOAC), 140 Metal-specific allergic contact dermatitis (ACD), 141 Mixed bed ion exchanger resin, 118 Multidimensional liquid chromatography (multi-LC), 69 Multiple Affinity Removal Spin Cartridge, 91 Multiple Affinity Removal System column, 30 Murashige and Skoog (MS) medium, 174 N Nanoflow reversed-phase liquid chromatography (nanoRPLC), 133 Nasal lavage fluid (NLF) analysis data interpretation, 84 by liquid chromatography and mass spectrometry, 79, 81–83 preparation, 80–81 acid-ethanol precipitation, 81 protein digestion, 81 total protein assay, 80 total ion chromatogram (TIC) profile of, 82

337 Nasal secretions nasal provocation, 79–80 pharmacological treatment, 78–79 preparation for proteome analysis, 77–78 Ni-affinity enriched proteins from human antigen, 141 Nickel-nitrilotriacetic acid (Ni-NTA), 142 Ni-NTA Magnetic Agarose Beads, 143 Nitrilotriacetic acid (NTA), 140 Nonequilibrium pH gradient gel electrophoresis (NEPHGE), 113 P Pancreatic adenocarcinoma cells proteome analysis, 4–5 Pancreatic intraepithelial neoplasia, 1–2 PanIN, Pancreatic intraepithelial neoplasia, 2 Parasitic protozoa cell fractionation, 313–314 cell culture and standard equipment, 315–316 electron microscopy, 316 hydrogenosome isolation and subfractionation, 327–329 isolation and subfractionation flagellum of trypanosomatids, 316 hydrogenosomes, 317 isolation of golgi complex, 316–317, 323–327 reservosome, 316, 320–321 trypanosomatid structures, 317–320 Pepper seed DNA, 161–162 Peptide mass fingerprinting (PMF), 12 Percoll gradient solutions, 165 Phenylmethylsulfonyl fluoride (PMSF), 116 PIR BLAST software, 84 Plant cell culture isolation of mitochondria, 163–169 crude organelle pellet differential centrifugation of, 165–166

338 density gradient purification of mitochondria, 167 materials, 164–165 protoplasts disruption and isolation, 165 Plant cell wall poteins isolation, 191–192 extraction and analysis from cell suspension cultures, 190–191 labile/weakly extraction by nondestructive techniques, 189–190 Rosette leaves extraction and analysis, 191–192 Plant material high-throughput extraction device DNA electrophoresis, 156, 159–160 inter simple sequence repeat PCR (ISSR-PCR), 156, 159 isoelectric focusing (IEF), 155–156, 158–159 sample preparation, 154–155, 157–158 Plant mitochondria Percoll gradient purification, 166 Plant proteases, 149 Plastids organelles, 171 Plastoglobuli lipid-containing structures, 173 ® POROS beads, 17 Protean IEF Cell, 70 Protease inhibitor cocktail tablets, 70 Protein concentration estimation kit, 176 content after, sinusitis NLFs pre- and post-pharmacological treatment, 82 deglycosylation, 281–282 with PNGase F, 282 trifluoromethanesulfonic acid with, 282–283 Proteins analysis by fluorescence dye saturation labeling and 2-DE, 2

Index cysteine-specific protein labeling using CyDye DIGE fluor saturation dyes, 2 2D-PAGE, 3 fluorescence dye labeling optimisation of, 6–7 isoelectric focusing, 2–3 microdissection and sample preparation, 2 microdissection for proteome analysis of pancreatic adenocarcinoma cells, 4 minimal number of microdissected cells determination of, 4–6 protein spots micropreparation of, 12 reference proteome for internal standardization and protein identification, 7–8 two-dimensional gel electrophoresis, 9–10 extraction from green plant tissue, 149–150, 152 materials and methods, 151 G POROS column, 20 G Sepharose, 58 identification and reference proteome for internal standardization, 8–9 loads for silver and Coomassie blue staining, 108 precipitation chloroform/methanol/trichloroacetic acid with, 280–281 quantitation by BCA Assay, 117–118 secretion from yeast protoplasts in active cell wall regeneration, 241 alkaline phosphatase assay, 247, 250–254 collection of proteins, 245–249 concentration of proteins secreted from regenerating protoplasts, 247, 250

Index preparation of yeast protoplasts, 249 recovery of proteins secreted from regenerating protoplasts, 246, 250 Protoplast formation efficiency of evaluation, 259 R Recombinant His-tagged proteins, 140 Reverse-phase C18 column, 79 Rosette leaves labile CWP analysis by 2-DE, 196 proteins extraction, 194–196 vacuum-infiltration, 195 S S. cerevisiae and C. albicans, cell wall, 218, 221–222 S. pyogenes strain 5448 cell wall proteome, 305 SEQUEST, 64 Silver staining procedures, 60 Size exclusion chromatography (SEC), 91 Spectra/Por® cellulose ester, 190 Speed Vac centrifugation, 73 Streptococcus mutans, silver stained 2-D gel, 290 T TCA/Acetone precipitation, 50, 120 Terminator device, 154–155 Triacyglycerol (TAG) biosynthesis, 203 Trichomonad structures isolation, 325 Trifluoromethanesulfonic acid, chemical protein deglycosylation with, 281–282 T ritrichomonas foetus, 315 T rypanosoma cruzi, 314 Trypsin-mediated incorporation of O18 , 137 Trypticase Yeast Maltose Medium (TYM), 315

339 U Ultraflex MALDI-TOF spectrometer, 143 Urinary protein concentrates, size exclusion chromatography, 90, 93 Urine samples preparation for proteomic analysis, 89 final sample preparation for proteomic analysis, 91 immunoaffinity subtraction of proteins, 91 size exclusion chromatography of urinary protein concentrates, 90, 92–94 urinary protein concentrate preparation, 90, 91

V Vivapure Anti-HSA/IgG removal kit, 91 Vmax® microplate reader, 117 Voyager DE-Pro mass spectrometer, 63

W Weakly bound CWP analysis, 191–192, 196 Weakly bound CWP extraction destructive techniques, 192, 196–197 cell wall preparation, 192, 196–197 protein extraction and separation, 192, 197 Western blot assembly, 106

Y Yeast and fungal cell walls protein solubilization -1,3 glucanase treatment, 224–225, 229 detergents and reducing agents, 224, 228

340 and exochitinase treatment, 225, 229–230 under mild alkali conditions, 224, 228–229 Yeast and fungal proteomics cell wall fractionation, 217–237 cell wall isolation from yeasts and filamentous fungi, 223–228

Index protein precipitation, 225, 230 protein solubilization from isolated yeast and fungal cell walls, 224–225, 228 Yeast proteome 2-D electrophoresis, 269–270 Yeast-Peptone-D-glucose (YPD), 222